18O稳定同位素标记定量蛋白质组研究技术的建立与优化

建立了18O稳定同位素标记方法,用于复杂体系蛋白质相对定量分析。对影响蛋白质标记稳定性的实验条件进行了比较和优化。结果表明,采用酶切后标记的方法,酶切肽段在胰酶催化下,在pH 5.0的K2HPO4/KH2PO4缓冲体系中,37℃18O标记反应16 h,绝大部分肽段即可达到100%的标记效率。对多个16O/18O成对肽段峰强度的动态范围及定量准确度进行了考察。结果表明,18O标记方法是一种简便、稳定、可靠的相对定量方法,10倍动态范围内,标记率相对标准偏差在18.4%以内,16O/18O峰强度呈很好的线性关系。本实验考察了标记后的肽段在不同溶液体系中的稳定性,为复杂样品的预处理和预分离的溶液条件提供了依据。     

18O同位素标记定量肽段串联体蛋白质结合同位素稀释-多反应监测质谱的蛋白质绝对定量新方法

建立了定量肽段串联体蛋白质(concatamers of Q peptides, QconCATs)结合¹⁸O同位素标记-多反应监测质谱的蛋白质绝对定量新方法。首先对QconCAT重组蛋白质进行了纯度表征,十二烷基硫酸钠-聚丙烯酰胺凝胶电泳(SDS-PAGE)表征结果表明重组蛋白质的纯度在99%以上,相对分子质量约为63.4 kDa。对QconCAT重组蛋白质酶切后的肽段混合物进行质谱分析,并经pFind和pLabel软件处理,验证了目标肽段。还考察了QconCAT重组蛋白质的酶切效率和¹⁸O标记效率,并对QconCAT蛋白质结合¹⁸O标记-同位素稀释-多反应监测质谱方法进行了评价。实验结果表明,采用该方法对腾冲嗜热厌氧菌(Thermoanaerobacter tengcongensis, TTE)中选定蛋白质的肽段进行绝对含量测定时,相对标准偏差小于20%,准确度较高,说明该方法可用于复杂生物样本中蛋白质的绝对定量。更重要的是所建方法不仅解决了细胞培养氨基酸稳定同位素标记(SILAC)技术的重标试剂价格昂贵的问题,也为定量蛋白质组学提供了一种新的方法。     

等电聚焦分离与18O稳定同位素标记联用的磷酸化蛋白质组学定量方法研究

磷酸化蛋白质组学定量分析,要对磷酸化修饰富集技术和定量技术进行研究。基于此,本研究采用18O稳定同位素标记技术对胰蛋白酶酶解肽段混合物进行标记,并对其标记时间和标记后胰蛋白酶的变性条件进行优化。结果表明:在pH=4～5的KH2PO4缓冲体系中,37℃,标记反应持续19～24h,除了C-端肽之外,几乎所有的肽段都可达到100%标记;采用TCEP可以有效地抑制16O-18O回标现象。建立了与18O标记技术兼容性良好的IPG-IEF技术对磷酸化肽段进行选择性富集,富集后共从HepG2细胞中鉴定到491个磷酸化位点、362个磷酸化肽段和356个磷酸化蛋白,表明IPG-IEF在大规模磷酸化肽段分离富集中是有效的;最后与高准确度高灵敏度高分辨率的LTQ-FTICR质谱仪联用,建立了基于18O-IPG-IEF-LTQ-FTICR的磷酸化蛋白质组定量技术。实验结果表明,该技术可以实现磷酸化肽段的有效定性和定量。本研究为磷酸化蛋白质组学定量研究提供了实用技术。     

基于末端准等重同位素标记的肽段从头测序方法

蛋白质的从头测序在蛋白质生物功能的研究、疾病相关蛋白突变体的发现和抗体药物表征等领域具有关键作用,其主要方法是将蛋白质酶解成肽段,通过二级谱图中碎片离子的质量差异判断氨基酸残基的序列信息,进而实现肽段的从头测序。由于谱图中的碎片离子信号易受噪声和不能用于测序的离子的干扰,目前的从头测序算法准确度较低。本研究发展了一种基于肽段末端准等重同位素标记(Pseudo-isobaric peptide termini labeling,PIPTL)的从头测序方法,将肽段样品分成两份,对一份肽段使用甲醛进行N端二甲基化标记,使用H₂¹⁸O进行C端¹⁸O标记;对另一份样品使用氘代甲醛进行N端二甲基化标记,C端不做处理;两份样品等量混合后,相同序列的不同标记的肽段质量仅相差0.016 Da,这些准等重肽段可在质谱中同时碎裂,产生成对的碎片离子。通过发展基于碎片离子成对性的测序算法简化谱图,并有效提取和分辨b/y离子,实现对肽段快速从头测序。本方法对牛血清白蛋白酶解后肽段测序准确度达到95.51%,显著优于商品化测序软件PEAKS的测序准...     

依赖色谱保留时间的智能化选择反应监测质谱方法的建立及其初步应用

建立了依赖色谱保留时间的智能化选择反应监测质谱方法,并与非依赖色谱保留时间的智能化选择反应监测质谱分析方法对不同体系(牛血清白蛋白酶切物、6种标准蛋白质混合物酶切物、腾冲嗜热菌蛋白提取液酶切物)的分析结果进行了系统比较。结果表明,引入色谱保留时间后的智能化选择反应监测质谱方法能够显著提高肽段及蛋白质的鉴定量,并且在复杂体系（如腾冲嗜热菌蛋白提取液酶切物）中效果尤为明显,鉴定到的肽段及蛋白质的覆盖率可分别达到目标肽段和蛋白质数量的89.62%和92.41%,并且灵敏度高、重复性好,能够实现对质荷比相同但保留时间有差异的肽段的准确鉴定。该方法将在复杂生物样本目标蛋白质组高通量、高灵敏度的鉴定、验证和确认中发挥独特作用。     

基于精氨酸酶切的蛋白质C端肽段富集方法的优化及评估北大核心CSCD

基于聚合物的蛋白质C端反向富集策略是用于研究蛋白质C端最为广泛的策略之一。目前,基于胰蛋白酶(trypsin)切割精氨酸残基C端(ArgC型酶切)的蛋白C端组学方法对蛋白质C端的鉴定深度仍有待提高。为解决这一问题,该研究对此方法进行了优化和评估:建立了基于“V型”过滤装置的“一锅法”富集流程,避免了副反应的干扰,缩短了样本的制备时间;优化了蛋白水平乙酰化反应条件,最大限度地降低了丝氨酸、苏氨酸、酪氨酸残基上的副反应,提高了肽段鉴定的可信性;优化了基于固相萃取枪头膜片过滤柱(StageTip柱)的样品分离过程,使C端肽段的鉴定深度增加至原来的4倍。通过以上优化,按照肽段水平错误发现率(FDR)<0.01、离子分数(ion score)≥20,且C端带有乙醇胺修饰的数据筛选标准,从人HEK 293T细胞中共鉴定出696个蛋白质C端。若仅要求肽段水平FDR<0.01,鉴定数目进一步增加到933个,这是基于聚合物富集策略的蛋白质C端组学方法所得的最大数据集之一。探索了胰蛋白酶镜像酶(LysargiNase)切割精氨酸残基N端(ArgN型酶切)与不同肽段N端衍生化修饰组合对蛋白质C端鉴定数目和种类的影响,“LysargiNase酶切+肽段N端乙酰化”新策略在原有“胰蛋白酶酶切+肽段N端二甲基化”策略的基础上将鉴定蛋白质C端的种类提升了47%。综上,该研究通过对基于Arg型酶切的蛋白C端组学方法的优化,提升了C端肽段的鉴定深度,扩大了C端肽段鉴定的覆盖范围。该方法将有望成为系统性表征蛋白质C端的有力工具。     

高效液相色谱-串联质谱法结合稳定同位素标记肽段同时测定稻米及其制品中3种过敏蛋白质

基于稳定同位素标记特征肽段和液相色谱-质谱联用仪建立稻米及制品中3种过敏蛋白质的同时定量方法。稻米及制品样品经盐溶液提取,赖氨酰基内切酶(Lys-C)和胰蛋白酶依次水解,C18-SD柱净化后,采用纳升高效液相色谱-线性离子阱-静电场轨道阱(NanoLC-LTQ-Orbitrap)采集和Protein Discovery软件鉴定,NCBI和Uniprot数据库的基本局部搜索比对工具(BLAST)筛选验证,最终获得表征稻米及制品中α-淀粉酶/胰蛋白酶抑制剂类蛋白质(seed allergenic protein RAG2, RAG2)、乙二醛酶Ⅰ活性蛋白(glyoxalase Ⅰ)和α-球蛋白(19 kDa globulin)3种过敏蛋白质的特异性肽段。3个特异性肽段经液相色谱梯度洗脱,在Poroshell色谱柱上实现完全分离,由三重四极杆质谱仪分析。实验通过优化多反应监测(MRM)质谱参数,比较不同溶剂体系、水解酶种类和酶量等酶解条件,结合内标法定量,实现对稻米及制品中3种蛋白质的绝对定量。实验结果表明,当酶解溶剂中含1 g/L十二烷基硫酸钠,采用Lys-C和胰蛋白酶组合消化策略,可有效提高3种蛋白质的酶切效率至65.7%~97.3%。该方法在1~200 nmol/L范围内线性关系良好,相关系数均大于0.9972, 3种蛋白质的检出限和定量限分别为3 mg/kg和10 mg/kg。3种蛋白质在空白稻米制品基质中3个水平下的加标回收率为80.6%~103.7%,日间和日内精密度均小于11.5%。该方法稳定性好,检测灵敏度高,操作简便,在分析各类稻米及制品中3种过敏蛋白质含量具有广泛的应用前景。     

基于蛋白质组学的细胞色素P450和葡萄糖醛酸转移酶亚型绝对定量分析

利用蛋白质组学方法,将大鼠肝微粒体样品进行胰蛋白酶水解;再利用液相色谱-串联质谱法(LCMS/MS),采用多反应监测模式(MRM),通过测定蛋白质水解后产生的特征酶切肽段,实现同时对大鼠肝微粒体内药物代谢酶P450和UGT的绝对定量。本实验首先建立标准工作曲线,对肝微粒体样品中P450和UGT进行定量,在线性范围内,相关系数r>0.995,线性关系良好,定量限≤10 nmol/L;以合成的稳定同位素标记特征肽段作为内标,对UGT1A1进行定量分析。结果表明,同位素标记特征肽段与未标记肽段色谱行为与质谱响应一致,在基质溶液中同位素标记肽段线性关系良好,利用标准曲线法和稳定同位素稀释法测得UGT1A1含量分别为17.30和18.23 nmol/g,两种方法所得结果基本一致,但稳定同位素稀释法操作简便,更适用于复杂样品的高通量测定。     

基于准等重二甲基化标记的高通量蛋白质组选择性交叉标记装置

面对生物学及精准医学等领域多变量、大样本量的蛋白质组定量分析的需求,高通量的定量标记及分析已经成为近年来蛋白质组学方法发展的趋势。发展了一种基于准等重二甲基化标记策略的高通量肽段末端选择性交叉标记装置（pIDL-StageTip）,借助简单的装置及离心力,有效地增加了定量标记的通量,并保证了肽段末端两步标记反应时间的可控性及操作的简便性。通过优化酸性条件下NaBD₃CN与NaBH₃CN体系的标记条件,得到了标准蛋白质酶解产物100%的标记效率、95%以上的标记选择性;在人源蛋白质组复杂体系下,标记效率大于99%,标记选择性为100%。基于该装置的定量方法具有很高的定量准确度及精密度。该装置为实现高可操作性、高准确度、高通量的蛋白质组定量标记提供了一个可靠的解决方案。     

基于共价色谱分离的含半胱氨酸肽段富集策略

多维色谱分离、串联质谱分析技术已在蛋白质组研究中得到广泛应用。然而生物样品的蛋白质以及全酶切肽段具有高度的复杂性,这严重干扰了蛋白质高通量、规模化的分析。通过标签肽段富集进行样品预分离可以降低体系的复杂程度。本文建立了一种基于共价色谱技术选择性分离富集含半胱氨酸肽的方法,从而降低了样品体系的复杂程度。首先以牛血清白蛋白(BSA)的酶切肽段为模型,对富集条件进行了优化和考察,并在此基础上通过5种蛋白质酶切肽段混合物的富集对该方法进行了验证。结果证明此方法的重现性好,富集效率高,富集特异性好,能有效地富集鉴定含半胱氨酸肽段。所建立的方法在复杂体系的蛋白质组研究中具有广泛的应用前景,为复杂样品的蛋白质高通量、自动化、规模化鉴定和定量研究提供了实用技术。     

18O labeling: a tool for proteomics.

An evaluation of the proteolytic labeling and quantification of proteins for diagnostic purposes using trypsin and 18O-enriched H2O is presented. We demonstrate that comparative or relative quantitation can be performed effectively with this approach. We have developed a protocol that allows the conservation of the labeled peptides in natural abundance water without fear of back-exchange providing that pH is sufficiently low to quench the catalytic activity of trypsin, but not so low as to promote chemical back-exchange. Because the labeling efficiency depends on the nature of the peptide, a simple linear relationship between the relative 16O/18O digest buffer mixture content (x) and labeling efficiency (y) does not exist; rather it follows a probability based y = x(2) relationship. As such, the extent of peptide labeling using 16O/18O digest buffer mixture ratios may deviate significantly from that expected based on a linear relationship. The evaluation of the relative Ziptip efficiency indicated a loss in sample recovery as the peptide concentration was reduced using normal conditions, suggesting that there is a limit below which there are diminishing returns. In addition, the adsorptive losses due to Speedvac dry down and recovery indicated modest (20%) losses that may vary widely (0-50%) from peptide to peptide. The in-solution digestion efficiency of standard protein mixtures as a function of concentration revealed a linear decrease with decreasing concentration. This is consistent with enzyme kinetic effects and emphasizes a potential quantitation error that could arise when evaluating differential expression based on peptide detection. The results from our studies demonstrate the power of 18O labeling as an optimization tool for proteomics process development.     

A method for calculating <Superscript>16</Superscript>o/<Superscript>18</Superscript>o peptide ion ratios for the relative quantification of proteomes

A method is described for the identification and relative quantification of proteomes using accurate mass tags (AMT) generated by nLC-dual ESI-FT-ICR-MS on a 7T instrument in conjunction with stable isotope labeling using 16O/18O ratios. AMTs were used for putative peptide identification, followed by confirmation of peptide identity by tandem mass spectrometry. For a combined set of 58 tryptic peptides from bovine serum albumin (BSA) and human transferrin, a mean mass measurement accuracy of 1.9 ppm +/-0.94 ppm (CIM99%) was obtained. This subset of tryptic peptides was used to measure 16O/18O ratios of 0.36 +/- 0.09 (CIM99%) for BSA (micro = 0.33) and 1.48 +/- 0.47 (CIM99%) for transferrin (micro = 1.0) using a method for calculating 16O/18O ratios from overlapping isotopic multiplets arising from mixtures of 16O, 18O1, and 18O2 labeled C-termini. The model amino acid averagine was used to calculate a representative molecular formula for estimating and subtracting the contributions of naturally occurring isotopes solely as a function of peptide molecular weight. The method was tested against simulated composite 16O/18O spectra where peptide molecular weight, 16O/18O ratio, 18O1/18O2 ratios, and number of sulfur atoms were varied. Relative errors of 20% or less were incurred when the 16O/18O ratios were less than three, even for peptides where the number of sulfur atoms was over- or under-estimated. These data demonstrate that for biomarker discovery, it is advantageous to label the proteome representing the disease state with 18O; and the method is not sensitive to variations in 18O1/18O2 ratio. This approach allows a comprehensive differentiation of expression levels and tentative identification via AMTs, followed by targeted analysis of over- and under-expressed peptides using tandem mass spectrometry, for applications such as the discovery of disease biomarkers.     

Accelerated 18O-labeling in urinary proteomics

Proteolytic (18)O-labeling of peptides has been studied and optimized in order to improve the labeling efficiency and to accelerate the process without increasing the degree of incomplete labeling. Using peptides generated from tryptic digested bovine serum albumin (BSA) and cytochrome c as model proteins, it was shown that complete labeling was achieved after 2 h at pH 6. To increase the sample throughput in a bottom-up proteomic setup, tryptic digestion of proteins in-solution was replaced with tryptic digestion using immobilized trypsin. As a result, an integrated approach was made possible, where both digestion (pH 8) and (18)O/(16)O-labeling of the resulting peptides (pH 6) were done using immobilized trypsin beads. This simplified the sample handling and reduced the overall reaction time significantly: the setup enabled tryptic digestion and (18)O/(16)O-labeling without sample transfer steps within 3.5 h with average (18)O/(16)O-ratios of 0.96±0.13 in aqueous buffer. The initial results were confirmed with a more complex matrix, by spiking urine with the model proteins, yielding results comparable with the ratios obtained in buffer. Satisfying ratios were also achieved regarding urinary proteins identified in a full scale bottom-up experiment. Average (18)O/(16)O-peptide ratios of 0.83±0.13 and 0.91±0.27 indicated good performance in a highly relevant matrix for biomarker discovery.     

Considerations for proteolytic labeling-optimization of 18O incorporation and prohibition of back-exchange

Proteolytic (18)O labeling is a very powerful tool for differential analysis applied to proteome studies. However, it is a relatively new technique and the optimization of the labeling process still needs some attention. We found that the two-step post-proteolytic labeling should be favored over the conventional digestion of proteins in H(2) (18)O, since the former allows for higher sample concentrations and thus more favorable kinetics. It was demonstrated that the inhibitory effect of urea on (18)O incorporation could be compensated by the use of higher sample concentrations. Furthermore, it was shown that heat-deactivation of trypsin prevents (18)O/(16)O back-exchange. In addition, no non-specific hydrolysis of the peptides could be observed as a result of the heating. Heat inactivation of trypsin opens the way for the use of capillary electrophoresis as a separation technique in proteolytic labeling studies, as it abolishes the need for use of detrimental additives. Analysis of a labeled protein digest by capillary isoelectric focusing/mass spectrometry showed the applicability of the method. No back-exchange was observed across the entire electropherogram.     

Use of 18O labels to monitor deamidation during protein and peptide sample processing

Nonenzymatic deamidation of asparagine residues in proteins generates aspartyl (Asp) and isoaspartyl (isoAsp) residues via a succinimide intermediate in a neutral or basic environment. Electron capture dissociation (ECD) can differentiate and quantify the relative abundance of these isomeric products in the deamidated proteins. This method requires the proteins to be digested, usually by trypsin, into peptides that are amenable to ECD. ECD of these peptides can produce diagnostic ions for each isomer; the c. + 58 and z - 57 fragment ions for the isoAsp residue and the fragment ion ((M + nH)((n-1)+.) - 60) corresponding to the side-chain loss from the Asp residue. However, deamidation can also occur as an artifact during sample preparation, particularly when using typical tryptic digestion protocols. With 18O labeling, it is possible to differentiate deamidation occurring during trypsin digestion which causes a +3 Da (18O1 + 1D) mass shift from the pre-existing deamidation, which leads to a +1-Da mass shift. This paper demonstrates the use of (18)O labeling to monitor three rapidly deamidating peptides released from proteins (calmodulin, ribonuclease A, and lysozyme) during the time course of trypsin digestion processes, and shows that the fast (approximately 4 h) trypsin digestion process generates no additional detectable peptide deamidations.     

Indirect ultrasonication for protein quantification and peptide mass mapping through mass spectrometry-based techniques

We report in this work a fast protocol for protein quantification and for peptide mass mapping that rely on ¹⁸O isotopic labeling through the decoupling procedure. It is demonstrated that the purity and source of trypsin do not compromise the labeling degree and efficiency of the decoupled labeling reaction, and that the pH of the labeling reaction is a critical factor to obtain a significant ¹⁸O double labeling. We also show that the same calibration curve can be used for MALDI protein quantification during several days maintaining a reasonable accuracy, thus simplifying the handling of the quantification process. In addition we demonstrate that ¹⁸O isotopic labeling through the decoupling procedure can be successfully used to elaborate peptide mass maps. BSA was successfully quantified using the same calibration curve in different days and plasma from a freshwater fish, Cyprinus carpio, was used to elaborate the peptide mass maps.     

A New Strategy of Using O18-Labeled Iodoacetic Acid for Mass Spectrometry-Based Protein Quantitation

A new O(18) labeling protocol is designed to assist quantitation of cysteine-containing proteins using LC/MS. Unlike other O(18) labeling strategies, the labeling is carried out at the intact protein level (prior to its digestion) during reduction/alkylation of cysteine side chains using O(18)-labeled iodoacetic acid (IAA). The latter can be easily prepared by exchanging carboxylic oxygen atoms of commercially available IAA in O(18)-enriched water at low pH. Since incorporation of the O(18) label in the protein occurs at the whole protein, rather than peptide level, the quantitation results are not peptide-dependent. The excellent stability of the label in mild pH conditions provides flexibility and robustness needed of sample processing steps following the labeling. In contrast to generally costly isotope labeling reagents, this approach uses only two relatively inexpensive commercially available reagents (IAA and H(2)O(18)). The feasibility of the new method is demonstrated using an 80?kDa human serum transferrin (hTf) as a model, where linear quantitation is achieved across a dynamic range spanning three orders of magnitude. The new approach can be used in quantitative proteomics applications and is particularly suitable for a variety of tasks in the biopharmaceutical sector, ranging from pharmacokinetic studies to quality control of protein therapeutics.     

Simultaneous quantification and identification using <Superscript>18</Superscript>O labeling with an ion trap mass spectrometer and the analysis software application “ZoomQuant”

Stable isotope labeling with (18)O is a promising technique for obtaining both qualitative and quantitative information from a single differential protein expression experiment. The small 4 Da mass shift produced by incorporation of two molecules of (18)O, and the lack of available methods for automated quantification of large data sets has limited the use of this approach with electrospray ionization-ion trap (ESI-IT) mass spectrometers. In this paper, we describe a method of acquiring ESI-IT mass spectrometric data that provides accurate calculation of relative ratios of peptides that have been differentially labeled using(18)O. The method utilizes zoom scans to provide high resolution data. This allows for accurate calculation of (18)O/(16)O ratios for peptides even when as much as 50% of a (18)O labeled peptide is present as the singly labeled species. The use of zoom scan data also provides sufficient resolution for calculating accurate ratios for peptides of +3 and lower charge states. Sequence coverage is comparable to that obtained with data acquisition modes that use only MS and MS/MS scans. We have employed a newly developed analysis software tool, ZoomQuant, which allows for the automated analysis of large data sets. We show that the combination of zoom scan data acquisition and analysis using ZoomQuant provides calculation of isotopic ratios accurate to approximately 21%. This compares well with data produced from (18)O labeling experiments using time of flight (TOF) and Fourier transform-ion cyclotron resonance (FT-ICR) MS instruments.     

Trypsin catalyzed 16O-to-18O exchange for comparative proteomics: tandem mass spectrometry comparison using MALDI-TOF,ESI-QTOF,and ESI-ion trap mass spectrometers

Quantitative or comparative proteome analysis was initially performed with 2-dimensional gel electrophoresis with the inherent disadvantages of being biased towards certain proteins and being labor intensive. Alternative mass spectrometry-based approaches in conjunction with gel-free protein/peptide separation have been developed in recent years using various stable isotope labeling techniques. Common to all these techniques is the incorporation, biosynthetically or chemically, of a labeling moiety having either a natural isotope distribution of hydrogen, carbon, oxygen, or nitrogen (light form) or being enriched with heavy isotopes like deuterium, (13)C, (18)O, or (15)N, respectively. By mixing equal amounts of a control sample possessing for instance the light form of the label with a heavy-labeled case sample, differentially labeled peptides are detected by mass spectrometric methods and their intensities serve as a means for direct relative protein quantification. While each of the different labeling methods has its advantages and disadvantages, the endoprotease (16)O-to-(18)O catalyzed oxygen exchange at the C-terminal carboxylic acid is extremely promising because of the specificity assured by the enzymatic reaction and the labeling of essentially every protease-derived peptide. We show here that this methodology is applicable to complex biological samples such as a subfraction of human plasma. Furthermore, despite the relatively small mass difference of 4 Da between the two labeled forms, corresponding to the exchange of two oxygen atoms by two (18)O isotopes, it is possible to quantify differentially labeled proteins on an ion trap mass spectrometer with a mass resolution of about 2000 in automated data dependent LC-MS/MS acquisition mode. Post column sample deposition on a MALDI target parallel to on-line ESI-MS/MS enables the analysis of the same compounds by means of ESI- and MALDI-MS/MS. This has the potential to increase the confidence in the quantification results as well as to increase the sequence coverage of potentially interesting proteins by complementary peptide ionization techniques. Additionally the paired y-ion signals in tandem mass spectra of (16)O/(18)O-labeled peptide pairs provide a means to confirm automatic protein identification results or even to assist de novo sequencing of yet unknown proteins.     

Characterization of differently processed forms of enolase 2 from Saccharomyces cerevisiae by two-dimensional gel electrophoresis and mass spectrometry

Two-dimensional gel electrophoresis, bioinformatics, and mass spectrometry are key analysis tools in proteome analysis. The further characterization of post-translational modifications in gel-separated proteins relies fully on data obtained by mass spectrometric analysis. In this study, stress-induced changes in protein expression in Saccharomyces serevisiae were investigated. A total of eleven spots on a silver-stained two-dimensional (2-D) gel were identified by matrix-assisted laser desorption/ionization (MALDI) peptide mass mapping to represent C and/or N-terminal processed forms of enolase 2. The processing sites were determined by MALDI peptide mass mapping using a variety of proteolytic enzymes, by optimizing the sample preparation procedure and by specific labeling of all C-termini derived from in-gel digestion using a buffer containing 16O:18O (1:1). Out of eleven processed forms of enolase 2, six were fully characterized and the approximate processing sites identified for the remaining five.