Molecular Vision 2011; 17:186-198 <>
Received 25 November 2010 | Accepted 14 January 2011 | Published 19 January 2011

Phosphoproteomics characterization of novel phosphorylated sites of lens proteins from normal and cataractous human eye lenses

Chun-Hao Huang,1,2 Yi-Ting Wang,3,4,5 Chia-Feng Tsai,3,6 Yu-Ju Chen,3,5,6 Jiahn-Shing Lee,7 Shyh-Horng Chiou1,2

The first three authors contributed equally to this work

1Graduate Institute of Medicine and Center for Research Resources and Development, Kaohsiung Medical University, Kaohsiung, Taiwan; 2Institute of Biological Chemistry, Academia Sinica, Taipei, Taiwan; 3Institute of Chemistry, Academia Sinica, Taipei, Taiwan; 4Chemical Biology and Molecular Biophysics Program, Taiwan International Graduate Program, Institute of Chemistry, Academia Sinica, Taipei, Taiwan; 5Institute of Biochemical Sciences, National Taiwan University, Taipei, Taiwan; 6Department of Chemistry, National Taiwan University, Taipei, Taiwan; 7Department of Ophthalmology, Chang-Gung Memorial Hospital, Chang-Gung University, Taipei, Taiwan

Correspondence to: Shyh-Horng Chiou, Center for Research Resources and Development, Kaohsiung Medical University, Kaohsiung 807 or Institute of Biological Chemistry, Academia Sinica, Taipei 115, Taiwan; Phone: (886)-7-3133874; FAX: (886)-7-3133434; email:


Purpose: Post-translational modification (PTM) of lens proteins is believed to play various roles in age-related lens function and development. Among the different types of PTM, phosphorylation is most noteworthy to play a major role in the regulation of various biosignaling pathways in relation to metabolic processes and cellular functions. The present study reported the quantitative analysis of the in vivo phosphoproteomics profiles of human normal and cataractous lenses with the aim of identifying specific phosphorylation sites which may provide insights into the physiologic significance of phosphorylation in relation to cataract formation.

Methods: To improve detection sensitivity of low abundant proteins, we first adopted SDS-gel electrophoresis fractionation of lens extracts to identify and compare the protein compositions between normal and cataractous lenses, followed by tryptic digestion, enrichment of phosphopeptides by immobilized metal affinity chromatography (IMAC) and nano-liquid chromatography coupled tandem mass spectrometry (nanoLC-MS/MS) analysis.

Results: By comprehensively screening of the phosphoproteome in normal and cataractous lenses, we identified 32 phosphoproteins and 73 phosphorylated sites. The most abundantly phosphorylated proteins are two subunits of β-crystallin, i.e., βB1-crystallin (12%) and βB2-crystallin (12%). Moreover, serine was found to be the most abundantly phosphorylated residue (72%) in comparison with threonine (24%) and tyrosine (4%) in the lens phosphoproteome. The quantitative analysis revealed significant and distinct changes of 19 phosphoproteins corresponding to 28 phosphorylated sites between these two types of human lenses, including 20 newly discovered novel phosphorylation sites on lens proteins.

Conclusions: The shotgun phosphoproteomics approach to characterize protein phosphorylation may be adapted and extended to the comprehensive analysis of other types of post-translational modification of lens proteins in vivo. The identification of these novel phosphorylation sites in lens proteins that showed differential expression in the cataractous lens may bear some unknown physiologic significance and provide insights into phosphorylation-related human eye diseases, which warrant further investigation in the future.


Human eye lenses are composed of elongated fiber cells, in which about 90% of total soluble proteins belong to three major classes of proteins, i.e., α-, β- and γ-crystallins [1,2]. Essentially these crystallins can exist in the eye lens with little turnover throughout the entire human lifespan albeit with various degrees of post-translational modification (PTM). Various types of PTM have been identified in animal eye lenses including especially human lenses, e.g.: 1. Deamidation [3,4], 2. Non-enzymatic glycosylation or glycation [5,6], 3. Oxidation of some amino acid residues of lens proteins such as tryptophan and methionine [7,8], 4. Sulfhydryl-disulfide oxidation [9,10], 5. Acetylation of NH2-terminal and lysine residues [11,12], 6. Truncation of crystallins [13,14], and 7. Phosphorylation [15-23]. Among these, phosphorylation is most noteworthy to play a major role in the regulation of various biosignaling pathways in relation to metabolic processes and cellular functions [24-26], which may include cancer development, aging, and cataract formation. Therefore, identification of protein phosphorylation and its exact phosphorylated residues in proteins or enzymes of interest are always considered as a preeminent and nontrivial task in the conventional structural and functional study of various cellular proteins. Mainly attributable to the recent advent and state-of-the-art instrumentation of proteomics, the investigation of protein phosphorylation has gradually become more amendable to routine analysis.

The recent explosion in available genomic sequence information is providing a useful sequence infrastructure for proteomics database. A major aspect of various proteomics strategies is the determination of protein identity (Protein ID) using analytical ‘‘fingerprints’’ or peptide mass fingerprinting (PMF) generated by digestion of proteins with specific enzymes such as trypsin, from which tandem mass (MS/MS) spectra of peptide fragments can then be used for comparison and confirmation of protein ID in available sequence databanks. The strategy based on the direct analysis of peptides generated from protein digestion by high-resolution liquid chromatographies coupled with tandem MS/MS spectrometry has facilitated the so-called “shotgun proteomics” for the identification of protein mixtures from any tissues of interest. Various MS/MS spectra can be algorithmically compared with predicted peptide spectra from sequence databases to identify the respective proteins. By combining with the recent development of capillary multidimensional liquid chromatography (capillary-MDLC), this shotgun proteomics approach is capable of characterizing proteins directly from entire cell lysates [27-31]. In shotgun proteomics, MDLC is a necessity to reduce sample complexity and increase dynamic range of protein identification. Recently mass spectrometric methods are being developed along the line that not only identifies proteins in a mixture but also compares the relative levels of protein expression between two different samples, i.e., quantitative shotgun proteomics.

The serious drawback of conventional gel-based 2-D gel proteomics lies in low sensitivity and under-representation for some special classes of proteins such as the extremely basic or acidic groups of proteins and membrane proteins [32-34]. In our previous study [35], phosphorylated peptides from trypsin-digested total protein mixtures of porcine lenses were concentrated and enriched on IMAC followed by identification of phosphopeptides on μLC-MS/MS. Gel-free IMAC phosphopeptide enrichment coupled with μLC-MS/MS analysis was found to be capable of identifying phosphorylated sites of various proteins from the whole lens extract. In this study, we have further applied quantitative shotgun proteomics to study and compare protein phosphorylation between normal and cataractous lens extracts to provide some basis to probe the role of phosphorylation in relation to cataract formation in vivo.


Materials and biologic tissues

Normal (30-year-old) and cataractous (68-year-old, Grade III of nuclear sclerosis) human lenses were obtained post mortem from the Department of Ophthalmology, Chang Gung Memorial Hospital, Taipei, Taiwan (J.-S. Lee). Eye lenses were kept and stored at −80 °C freezer before dissection. Triethylammonium bicarbonate (TEABC) and iron chloride (FeCl3) were purchased from Sigma Aldrich (St. Louis, MO). The BCATM protein-assay reagent kit was obtained from Pierce (Rockford, IL). Ammonium persulfate and N,N,N’,N’-tetramethylenediamine were purchased from Amersham Pharmacia (Piscataway, NJ). Reagent-grade acetic acid (AA) was purchased from J. T. Baker (Phillipsburg, NJ). Trifluoroacetic acid (TFA), formic acid (FA) and HPLC-grade acetonitrile were purchased from Sigma Aldrich. Chemically-modified and sequencing-grade trypsin was purchased from Promega (Madison, WI).

Preparation of lens extracts

Lenses were homogenized and suspended in 20 mM Tris-HCl, pH 6.8 buffer containing 0.1% SDS and centrifuged for 30 min at 20,000× g for the extraction of total lens proteins as described previously [36-38].

1-D gel SDS–PAGE

After estimation of protein content by using a BCATM protein-assay reagent kit (Pierce, Rockford, IL), 10 μg of proteins in lens extracts were loaded on 12.5% one-dimensional SDS–PAGE for protein separation, followed by staining with Coomassie brilliant blue R-250 and destained in 10% methanol/ 7% acetic acid.

In-gel digestion and nanoLC-ESI-MS/MS

Based on the SDS–PAGE analysis of samples, differentially expressed proteins were selected for further identification by nanoLC-MS/MS. The protein bands separated on 1-D SDS–PAGE were cut from gels, and then destained three times with 25 mM ammonium bicarbonate buffer (pH 8.0) in 50% acetonitrile (ACN) for 1 h. The gel pieces were dehydrated in 100% ACN for 5 min and then dried for 30 min in a vacuum centrifuge. Enzyme digestion was performed by adding 0.5 μg trypsin in 25 mM ammonium bicarbonate buffer per sample at 37 °C for 16 h. The peptide fragments were extracted twice with 50 μl 50% ACN/ 0.1% TFA. After removal of ACN and TFA by centrifugation in a vacuum centrifuge, samples were dissolved in 0.1% formic acid/ 50% ACN and analyzed by nanoLC-ESI-MS/MS at the core facility laboratory of the Center for Research Resources and Development, Kaohsiung Kaohsiung Medical University, Kaohsiung, Taiwan and at Institute of Chemistry, Academia Sinica, Taipei, Taiwan. Proteins were identified in the NCBI databases by use of MS/MS ion search with the search program MASCOT as described previously [35].

Gel-assisted digestion

The protein samples from lenses were subjected to gel-assisted digestion. The sample was incorporated into a gel directly in an Eppendorf vial with acrylamide/bisacrylamide solution (40%, v/v, 29:1), 10% (w/v) APS, 100% TEMED in a proportion of 14:5:0.7:0.3. The gel was cut into small pieces and washed several times with 25 mM TEABC containing 50% (v/v) ACN. The gel samples were further dehydrated with 100% ACN and completely dried using a SpeedVac (ASAHI TECHNO GLASS Corp., Tokyo, Japan). Proteolytic digestion was then performed with trypsin (protein: trypsin=50:1, w/w [g/g]) in 25 mM TEABC with incubation overnight at 37 °C. The tryptic peptides were dried completely under vacuum and stored at −30 °C.

IMAC preparation and protocol

This step of sample preparation and procedure is most critical for a successful phosphoproteomics study of complex protein mixtures isolated from biologic tissues. The IMAC column was first capped one end with a 0.5 μm frit disk enclosed in stainless steel column-end fitting. The Ni-nitrilotriacetic acid (Ni-NTA) resin was extracted from spin column (Qiagen, Hilden, Germany) and packed into a 10-cm microcolumn (500 μm i.d. PEEK column, Upchurch Scientific/Rheodyne, Oak Harbor, WA) as described previously [39]. Automatic purification of phosphopeptides was performed by connecting to an autosampler in an HP1100 solvent delivery system (Hewlett-Packard, Palo Alto, CA) with a flow rate 13 µl/min. First, the Ni2+ ions were removed with 100 µl 50 mM EDTA in 1 M NaCl. Then the IMAC column was activated with 100 µl 0.2 M FeCl3 and equilibrated with loading buffer for 30 min before sample loading. The loading buffer/ acetic acid was 6% (v/v) and the pH was adjusted to 3.0 with 0.1 M NaOH (pH=12.8). The peptide samples from trypsin digestion were reconstituted in the loading buffer and loaded into the IMAC column that had been equilibrated with the same loading buffer for 20 min. Then the unbound peptides were removed with 100 μl washing solution consisting of 75% (v/v) loading buffer and 25% (v/v) ACN, followed by equilibration with loading buffer for 15 min. Finally, the bound peptides were eluted with 100 µl 200 mM NH4H2PO4 (pH 4.4). Eluted peptide samples were dried under vacuum and then reconstituted in 0.1% (v/v) TFA (40 μl) for further desalting and concentration using ZipTipsTM (Millipore, Bedford, CA).

NanoLC-MS/MS analysis

Purified phosphopeptide samples from about 500 µg total protein extract were reconstituted in 4 µl buffer A (0.1% formic acid (FA) in H2O) and analyzed by LC-Q-TOF MS (Waters Q-TOFTM Premier; Waters Corp, Milford, MA). For LC-MS/MS analysis by Waters Q-TOFTM Premier system, samples were injected into a 2 cm×180 μm capillary trap column and separated by 20 cm×75 μm Waters1 ACQUITYTM 1.7 mm BEH C18 column using a nanoACQUITY Ultra Performance LCTM system (Waters Corp., Milford, MA). The column was maintained at 35 °C and bound peptides were eluted with a linear gradient of 0%–80% buffer B (buffer A, 0.1% FA in H2O; buffer B, 0.1% FA in ACN) for 120 min. MS was operated in ESI positive V mode with a resolving power of 10,000. NanoLockSpray source was used for accurate mass measurement and the lock mass channel was sampled every 30 s. The mass spectrometer was calibrated with a synthetic human [Glu1]-fibrinopeptide B solution (1 pmol/µl, from Sigma Aldrich) delivered through the NanoLockSpray source. Data acquisition was operated in the data directed analysis (DDA). The method included a full MS scan (m/z 400–1600, 0.6 s) and 3 MS/MS scans (m/z 100–1990, 1.2 s each scan) sequentially on the three most intense ions present in the full scan mass spectrum.

Database search and data processing/filtering

Raw MS/MS data were converted into peak lists using Distiller (version 2.0; Matrix Science, London, UK) with default parameters. All MS/MS samples were analyzed using Mascot (version 2.2.1; Matrix Science). Mascot was set up to search the Swissprot_Mammalia (version 54.2, 55307 entries) assuming trypsin as the digestion enzyme. MASCOT was searched with a fragment ion mass tolerance of 0.1 Da and a parent ion tolerance of 0.1 Da. Two missed cleavages were allowed for trypsin digestion. Phosphorylation (Ser/Thr/Tyr) and oxidation (Met) were selected as two variable modifications. To evaluate the false discovery rate of protein identification, we repeated the search using identical search parameters and validation criteria against a randomized decoy database created by MASCOT. The false discovery rates with MASCOT score >36 (p<0.05) was 0.73% in our phosphoproteomics study of lens protein extracts.

Label-free quantitation method

The quantitative analysis of peptides in the label-free experiments was performed by employing our recently published software, IDEAL-Q [40,41]. The raw data files acquired from Waters Q-TOFTM Premier were converted into files of mzXML format by the program massWolf, and the search results in MASCOT were exported in eXtensive Markup Language data (.XML) format. After data conversion, the confident peptide identification results (p<0.05) from each LC-MS/MS run were loaded and merged to establish a global peptide information list (sequence, elution time, and mass-to-charge). Alignment of elution time is then performed based on the peptide information list using linear regression in different LC-MS/MS runs followed by correction of aberrational chromatographic shift across fragmental elution-time domains. To calculate relative peptide abundance, the tool performs reconstruction of extracted ion chromatography (XIC), and calculation of XIC area. The fold-change of a given peptide was calculated by the ratio of relative peptide abundance between different samples.

Results and discussion

In spite of the biologic significance and physiologic role of protein phosphorylation and the rapid advances in MS methodologies, high-throughput characterization of site-specific phosphorylation residues in proteins is still challenged by the technical difficulties [42,43] associated with their dynamic modification patterns, substoichiometric concentrations, heterogeneous forms of phosphoproteins, and low sensitivity and response from MS analyses of total protein mixtures extracted from biologic tissues. Therefore improved methodologies that specifically enrich the transient phosphoproteome in a routine and comprehensive manner are important for studying phosphorylation-dependent cellular signaling associated with various diseased states [44].

Experimental design and methodology evaluation

Identification of large numbers of phosphopeptides with high specificity, reproducibility and recovery is critical in phosphoproteomics analysis. IMAC takes advantage of the phosphate groups as electron donors that chelate metal ion (Fe3+-NTA-silica) to preferentially retain phosphopeptides. Although the simple and routinely used protocol yields adequate results for simple phosphoprotein mixtures, the results for proteome-wide analysis are far from satisfactory. As shown previously [35,39], we have found the IMAC protocol used herein can yield an efficient enrichment and obtain specific purification for phosphopeptides devoid of contamination (a lack of nonspecific competitive binding). The pH effect for the binding and elution of phosphopeptides in IMAC protocol has been critically evaluated, demonstrating that the current IMAC method can reflect the representative phosphorylated amino-acid distribution such as phosphotyrosine, phosphoserine and phosphothreonine in the cell without bias. To date, the specificity and recovery reported in our IMAC protocol significantly exceed those previously achieved by single-step IMAC or IMAC in combination with methylation [45]. This protocol demonstrated high specificity (98%) that was comparable with TiO2 chromatography [46,47]. As compared to two-step purification methods, our protocol provides comparable selectivity and low sample loss with some advantages over current procedures. In terms of practical use, it offers a simple one-step, more reproducible method amenable to automatic phosphopeptide purification and enrichment using a Fe3+-IMAC microcolumn. Greater than 90% column recovery and enrichment specificity can be routinely achieved for single IMAC purification of up to 1 mg of protein lysates from various cell lines or tissues.

Gel-based 1D- or 2D-gel proteomics

In 1993, Henzel et al. [48] first reported and started the popular gel-based proteomics analysis by combining 2D-gel electrophoresis and mass spectrometry. The global identification of proteins in biologic samples was based on pre-separation of a protein mixture on 2-D gel electrophoresis. The mass spectral patterns from tandem mass (MS/MS) analysis of protein fragments generated from protease digestion were then compared with predicted peptide spectra from sequence databases to identify the respective proteins. Although previously 2-D gel electrophoresis coupled with tandem MS has been considered as the method of choice in proteomics study, only up to 2,000 individual polypeptide chains at most can be resolved on a single 2-D gel [31]. The number of detected proteins is still being relatively small as compared to the whole genome-encoded functional proteins of about 20,000~30,000 in higher vertebrates. It is especially under-representative of some special classes of proteins such as low-abundance transcription factors and membrane proteins [32-34] because of the low solubility of these classes of proteins in the first dimensional isoelectric-focusing (IEF) protein separation of 2-D gel electrophoresis in the absence of SDS denaturing agent. In our proteomic study of porcine lens proteins [35], we have also encountered poor solubility of some proteins in pre-MS 2-D gel separation.

To improve the detection sensitivity for low abundant proteins, fractionation was first performed for the total protein extracts of normal and cataractous human lenses by 1-D SDS–PAGE gels (Figure 1). The proteins were separated into at least more than 10 different protein bands or zones from total protein mixtures of normal and cataract lenses. In comparison with normal human lens-proteome, only four crystallin proteins, i.e., β-crystallin B1, β-crystallin B2, βs-crystallin B1, and γD-crystallin in cataractous lens showed significant decrease in their expression levels. Similar results in previous reports also pointed to the important role of differential crystallin expression leading to cataract formation [49-51]. It can be seen that 1-D gels are less tedious and time-consuming than 2-D gels and still afford a respectable and extensive protein separation capable of protein ID analysis after LC-MS/MS. The unambiguous identification of some major classes of β- and γ-crystallin classes were confirmed and verified in addition to α-crystallins reported previously [35]. However, similar to the previous 2-D gel phosphoproteomic study of α-crystallins, the phosphorylated sites identified by 1-D gel-based methodology are still very limited; only a few well known abundant and predominant sites such as Ser-59, Ser-81 and Ser-155 in αA-crystallin, Ser-19, Ser-21 and Ser-59 in αB-crystallin, and Thr-189, Ser-9 and Ser-95 in βB1-crystallin were identified [35]. We could not find any other phosphopeptides in trypsin-digested protein bands corresponding to other lens proteins when using the 1-D or 2-D gel approach probably due to lower abundance of phosphopeptides generated from digestion of protein bands. Therefore we have resorted to the newer strategy of quantitative shotgun proteomics by using IMAC for the enrichment of phosphopeptides generated from the protease digestion of total lens extracts.

Gel-free proteomic analysis of phosphorylated proteins in human lenses

Because the capability of a gel-based proteomic approach to identify phosphoproteins was limited for phosphopeptide identification, we adopted instead a gel-free protocol similar to shotgun proteomic approaches [28,29]. By enrichment of the lens phosphopeptides on IMAC followed by LC-MS/MS analysis, we have identified 73 phosphorylation sites in human lens proteins (Table 1). As shown in Table 1, the identified 172 nondegenerate phosphopeptides belonged to 32 proteins in the human lens, including 9 crystallin proteins and other non-crystallin lens proteins possessing different cellular functions. Among the identified phosphoproteins, the relative proportions of the corresponding proteins with functions relating to protein folding, metabolism and cytoskeleton were 32%, 28%, and 25%, respectively (Figure 2). The other 15% phosphoproteins consisted of proteins with specified functions of transport, cellular redox system and homeostasis.

As shown in Figure 3A, further analysis of the whole phosphoproteome in human lenses indicated that phosphorylation on serine (72%) was more common than that on threonine (24%) and tyrosine (4%). In Figure 3B, most phosphopeptides were identified as being crystallin proteins, indicating that the major classes of lens crystallins are also the most abundant phosphoproteins in the human lens tissue. The proportions of phosphopeptides identified as being βB1-crystallin, βB2-crystallin, αB-crystallin, γD-crystallin, filensin, αA-crystallin, and βs-crystallin were 12%, 12%, 9%, 8%, 8%, 6%, and 6%, respectively, emphasizing the fact that βB-crystallin subunits are indeed the major phosphorylation targets in the lens and may play a significant role in the phosphorylation-related biosignaling function in this transparent lens tissue.

Identification of phosphorylation sites in human lens crystallins

As shown in Table 1, the phosphorylation sites of crystallin proteins were found to spread over the entire polypeptide regions of these crystallins. Based on the proportion of phosphorylation sites in each crystallin, we found that Ser-81 (31%) and Ser93/Thr-118 (25%) are the predominant phosphorylation-sites in βB1- and βB2-crystallin, respectively (Figure 4A,B). In addition, the phosphorylation of αB-crystallin was shown to distribute evenly over the whole crystallin molecule at Ser-19 (23%), Ser-21 (22%), Ser-59 (22%), and Ser-139 (22%; Figure 4C) similar to our previous report on porcine αB-crystallin [35]. In contrast, some predominant phosphorylation sites present in other crystallin proteins were also identified, e.g., Ser-75 (50%) in γD-crystallin, Thr-148 (33%) in αA-crystallin, and Tyr-11/Ser-167 (33%) in β-crystallin S (also denoted as βs-crystallin). The mechanisms underlying the differential phosphorylation at specific sites of these crystallins remain unknown, which should be of interest for further study in the future.

Identification of phosphorylation sites in non-crystallin proteins involved in cytoskeleton, metabolism, transport, and cellular redox homeostasis

In addition to 9 lens crystallins, 23 non-crystallin proteins were also found to be phosphorylated in vivo in our phosphoproteomic analysis (Table 1). It is noteworthy that similar to αB-crystallin (a member of the small heat-shock protein family in the lens), another heat-shock protein beta-1 (homolog of heat-shock proteins Hsp27 and Hsp20) with chaperone activity was shown to be phosphorylated at Ser-82 [52,53]. To date these phosphorylated sites in the non-crystallin proteins have never been identified in the lens tissue and warranted for detailed functional characterization in the future.

Comparative analysis of phosphoproteome in human lenses with or without cataract

To investigate the differential post-translational modification of human lens proteins with or without cataract, we performed quantitative phosphoproteomic analysis. As shown in Table 2, 19 phosphoproteins consisting of 8 crystallin proteins and 11 non-crystallin proteins with their corresponding 28 phosphorylated sites were identified between these two types of human lenses. Among these identified proteins, the extents of 15 phosphorylated sites were found to increase by twofold while 13 sites decreased in phosphorylation, indicating that complicated post-translational modification such as phosphorylation may be one of the causative factors underlying the development of human cataract. Furthermore, some quantitative changes in the phosphorylation status were found even in the same proteins from normal and diseased lenses such as βB1-crystallin and αB-crystallin. In human cataractous lenses, two phosphorylated sites in βB1-crystallin, Ser-32 and Ser-81, were found to decrease while Ser-93 increased in their relative phosphorylation ratios (P/N ratio in Table 2). The different proportions of phosphorylation and specific phosphorylated sites associated with normal and cataractous human lens proteins may form a firm basis for unraveling the mechanistic pathways of cataract formation with aging. Furthermore, among the differentially expressed phosphopeptides, 20 phosphorylation sites were verified to be newly discovered based on comparison with those in the phosphoprotein databases, Uni-Prot and PhosphoSitePlus* website. The data also revealed 14 novel phosphorylation sites on 7 crystallin proteins. These differentially expressed phosphorylation and their associated phosphorylated sites might be the potential therapeutic targets of cataract disease, which warrant further investigation.


The conventional gel-based phosphoproteomics analyses by 1-D SDS–PAGE coupled LC-MS/MS and separately by IMAC enrichment of phosphopeptides followed by shotgun label-free quantitation method have been used to analyze and compare phosphorylation patterns of lens proteins from whole tissue extracts of normal and cataractous lenses. In this report we have focused on employing efficient IMAC protocol of phosphopeptide enrichment for profiling and quantitative analysis of transiently phosphorylated proteins. The IMAC protocol reported herein demonstrated enrichment with high specificity and low sample loss without the need for additional esterification and desalting step. This procedure may be applicable to a variety of materials such as tissue, cell and body fluid. As judged by the higher sample recovery and greater number of phosphopeptides identified by the critically validated IMAC procedure [35,39] in this study as compared to previous reports on phosphorylation analysis in the literature [4,15-18,54,55], it should prove feasible for the routine phosphoproteome analysis in the future. The combination of this protocol with either stable isotope tagging or a label-free methodology may be further employed for large-scale comparative proteomic studies to decipher the dynamic and complicated phosphoproteomes from various biologic samples of diverse tissues. On the other hand, the identification of these novel phosphorylation sites in lens proteins that showed differential expression in the cataractous lens may bear some as-yet-unknown physiologic significance and provide insights into phosphorylation-related human eye diseases.


This work was supported in part by Kaohsiung Medical University, Academia Sinica and the National Science Council (NSC Grants 96-2311-B-037-005-MY3, 99-2314-B-037-042 and 99-2745-B-037-005 to S.-H. Chiou), Taipei, Taiwan. Communication may be directed to Dr. Shyh-Horng Chiou and Dr. Yu-Ju Chen ( since they contributed equally to the research presented in this article and are to be considered co-corresponding authors.


  1. Harding JJ, Dilley KJ. Structural proteins of the mammalian lens: a review with emphasis on changes in development, aging and cataract. Exp Eye Res. 1976; 22:1-73. [PMID: 767125]
  2. de Jong WW, Hendriks W, Mulders JW, Bloemendal H. Evolution of eye lens crystallins: the stress connection. Trends Biochem Sci. 1989; 14:365-8. [PMID: 2688200]
  3. Van Kleef FS, De Jong WW, Hoenders HJ. Stepwise degradations and deamidation of the eye lens protein alpha-crystallin in ageing. Nature. 1975; 258:264-6. [PMID: 1202360]
  4. Miesbauer LR, Zhou X, Yang Z, Sun Y, Smith DL, Smith JB. Post-translational modifications of water-soluble human lens crystallins from young adults. J Biol Chem. 1994; 269:12494-502. [PMID: 8175657]
  5. Chiou SH, Chylack LT, , Jr Tung WH, Bunn HF. Nonenzymatic glycosylation of bovine lens crystallins. Effect of aging. J Biol Chem. 1981; 256:5176-80. [PMID: 7228874]
  6. Nagaraj RH, Sell DR, Prabhakaram M, Ortwerth BJ, Monnier VM. High correlation between pentosidine protein crosslinks and pigmentation implicates ascorbate oxidation in human lens senescence and cataractogenesis. Proc Natl Acad Sci USA. 1991; 88:10257-61. [PMID: 1946446]
  7. Andley UP, Clark BA. Generation of oxidants in the near-UV photooxidation of human lens alpha-crystallin. Invest Ophthalmol Vis Sci. 1989; 30:706-13. [PMID: 2703311]
  8. McDermott M, Chiesa R, Roberts JE, Dillon J. Photooxidation of specific residues in alpha-crystallin polypeptides. Biochemistry. 1991; 30:8653-60. [PMID: 1888728]
  9. Spector A. The search for a solution to senile cataracts. Proctor lecture. Invest Ophthalmol Vis Sci. 1984; 25:130-46. [PMID: 6321383]
  10. Hanson SR, Hasan A, Smith DL, Smith JB. The major in vivo modifications of the human water-insoluble lens crystallins are disulfide bonds, deamidation, methionine oxidation and backbone cleavage. Exp Eye Res. 2000; 71:195-207. [PMID: 10930324]
  11. Lapko VN, Smith DL, Smith JB. In vivo carbamylation and acetylation of water-soluble human lens alphaB-crystallin lysine 92. Protein Sci. 2001; 10:1130-6. [PMID: 11369851]
  12. Bloemendal H, de Jong W, Jaenicke R, Lubsen NH, Slingsby C, Tardieu A. Ageing and vision: structure, stability and function of lens crystallins. Prog Biophys Mol Biol. 2004; 86:407-85. [PMID: 15302206]
  13. Chaves JM, Srivastava K, Gupta R, Srivastava OP. Structural and functional roles of deamidation and/or truncation of N- or C-termini in human alpha A-crystallin. Biochemistry. 2008; 47:10069-83. [PMID: 18754677]
  14. Liao JH, Lee JS, Wu SH, Chiou SH. COOH-terminal truncations and site-directed mutations enhance thermostability and chaperone-like activity of porcine alphaB-crystallin. Mol Vis. 2009; 15:1429-44. [PMID: 19641632]
  15. Spector A, Chiesa R, Sredy J, Garner W. cAMP-dependent phosphorylation of bovine lens alpha-crystallin. Proc Natl Acad Sci USA. 1985; 82:4712-6. [PMID: 2991889]
  16. Chiesa R, Gawinowicz-Kolks MA, Kleiman NJ, Spector A. Definition and comparison of the phosphorylation sites of the A and B chains of bovine alpha-crystallin. Exp Eye Res. 1988; 46:199-208. [PMID: 3350065]
  17. Voorter CE, de Haard-Hoekman WA, Roersma ES, Meyer HE, Bloemendal H, de Jong WW. The in vivo phosphorylation sites of bovine alpha B-crystallin. FEBS Lett. 1989; 259:50-2. [PMID: 2599111]
  18. Smith JB, Thevenon-Emeric G, Smith DL, Green B. Elucidation of the primary structures of proteins by mass spectrometry. Anal Biochem. 1991; 193:118-24. [PMID: 2042736]
  19. Kamei A, Hamaguchi T, Matsuura N, Masuda K. Does post-translational modification influence chaperone-like activity of alpha-crystallin? I. Study on phosphorylation. Biol Pharm Bull. 2001; 24:96-9. [PMID: 11201254]
  20. Ito H, Kamei K, Iwamoto I, Inaguma Y, Nohara D, Kato K. Phosphorylation-induced change of the oligomerization state of alpha B-crystallin. J Biol Chem. 2001; 276:5346-52. [PMID: 11096101]
  21. Aquilina JA, Benesch JL, Ding LL, Yaron O, Horwitz J, Robinson CV. Phosphorylation of alphaB-crystallin alters chaperone function through loss of dimeric substructure. J Biol Chem. 2004; 279:28675-80. [PMID: 15117944]
  22. Ahmad MF, Raman B, Ramakrishna T, Rao Ch M. Effect of phosphorylation on alpha B-crystallin: differences in stability, subunit exchange and chaperone activity of homo and mixed oligomers of alpha B-crystallin and its phosphorylation-mimicking mutant. J Mol Biol. 2008; 375:1040-51. [PMID: 18061612]
  23. Aggeli IK, Beis I, Gaitanaki C. Oxidative stress and calpain inhibition induce alpha B-crystallin phosphorylation via p38-MAPK and calcium signalling pathways in H9c2 cells. Cell Signal. 2008; 20:1292-302. [PMID: 18420382]
  24. Pawson T, Scott JD. Signaling through scaffold, anchoring, and adaptor proteins. Science. 1997; 278:2075-80. [PMID: 9405336]
  25. Irvine R. Cell signaling. The art of the soluble. Science. 2007; 316:845-6. [PMID: 17495162]
  26. Petrini JH. Cell signaling. A touching response to damage. Science. 2007; 316:1138-9. [PMID: 17525326]
  27. Zhou H, Watts JD, Aebersold R. A systematic approach to the analysis of protein phosphorylation. Nat Biotechnol. 2001; 19:375-8. [PMID: 11283598]
  28. Washburn MP, Wolters D, Yates JR, 3rd. Large-scale analysis of the yeast proteome by multidimensional protein identification technology. Nat Biotechnol. 2001; 19:242-7. [PMID: 11231557]
  29. McCormack AL, Schieltz DM, Goode B, Yang S, Barnes G, Drubin D, Yates JR, 3rd. Direct analysis and identification of proteins in mixtures by LC/MS/MS and database searching at the low-femtomole level. Anal Chem. 1997; 69:767-76. [PMID: 9043199]
  30. MacCoss MJ, McDonald WH, Saraf A, Sadygov R, Clark JM, Tasto JJ, Gould KL, Wolters D, Washburn M, Weiss A, Clark JI, Yates JR, 3rd. Shotgun identification of protein modifications from protein complexes and lens tissue. Proc Natl Acad Sci USA. 2002; 99:7900-5. [PMID: 12060738]
  31. O'Farrell PH. High resolution two-dimensional electrophoresis of proteins. J Biol Chem. 1975; 250:4007-21. [PMID: 236308]
  32. Chiou S-H, Wu S-H. Evaluation of commonly used electrophoretic methods for the analysis of proteins and peptides and their application to biotechnology. Anal Chim Acta. 1999; 383:47-60.
  33. Han CL, Chien CW, Chen WC, Chen YR, Wu CP, Li H, Chen YJ. A multiplexed quantitative strategy for membrane proteomics: opportunities for mining therapeutic targets for autosomal dominant polycystic kidney disease. Mol Cell Proteomics. 2008; 7:1983-97. [PMID: 18490355]
  34. Leth-Larsen R, Lund RR, Ditzel HJ. Plasma membrane proteomics and its application in clinical cancer biomarker discovery. Mol Cell Proteomics. 2010; 9:1369-82. [PMID: 20382631]
  35. Chiou SH, Huang CH, Lee IL, Wang YT, Liu NY, Tsay YG, Chen YJ. Identification of in vivo phosphorylation sites of lens proteins from porcine eye lenses by a gel-free phosphoproteomics approach. Mol Vis. 2010; 16:294-302. [PMID: 20182557]
  36. Chiou SH, Azari P, Himmel ME, Squire PG. Isolation and physical characterization of bovine lens crystallins. Int J Pept Protein Res. 1979; 13:409-17. [PMID: 457334]
  37. Liao JH, Hung CC, Lee JS, Wu SH, Chiou SH. Characterization, cloning, and expression of porcine alpha B crystallin. Biochem Biophys Res Commun. 1998; 244:131-7. [PMID: 9514893]
  38. Lee JS, Samejima T, Liao JH, Wu SH, Chiou SH. Physiological role of the association complexes of alpha-crystallin and its substrates on the chaperone activity. Biochem Biophys Res Commun. 1998; 244:379-83. [PMID: 9514930]
  39. Tsai CF, Wang YT, Chen YR, Lai CY, Lin PY, Pan KT, Chen JY, Khoo KH, Chen YJ. Immobilized metal affinity chromatography revisited: pH/acid control toward high selectivity in phosphoproteomics. J Proteome Res. 2008; 7:4058-69. [PMID: 18707149]
  40. Tsou CC, Tsai CF, Tsui YH, Sudhir PR, Wang YT, Chen YJ, Chen JY, Sung TY, Hsu WL. IDEAL-Q, an automated tool for label-free quantitation analysis using an efficient peptide alignment approach and spectral data validation. Mol Cell Proteomics. 2010; 9:131-44. [PMID: 19752006]
  41. Wang YT, Tsai CF, Hong TC, Tsou CC, Lin PY, Pan SH, Hong TM, Yang PC, Sung TY, Hsu WL, Chen YJ. An informatics-assisted label-free quantitation strategy that depicts phosphoproteomic profiles in lung cancer cell invasion. J Proteome Res. 2010; 9:5582-97. [PMID: 20815410]
  42. Mann M, Ong SE, Gronborg M, Steen H, Jensen ON, Pandey A. Analysis of protein phosphorylation using mass spectrometry: deciphering the phosphoproteome. Trends Biotechnol. 2002; 20:261-8. [PMID: 12007495]
  43. Chen CH. Review of a current role of mass spectrometry for proteome research. Anal Chim Acta. 2008; 624:16-36. [PMID: 18706308]
  44. Hunter T. Protein kinases and phosphatases: the yin and yang of protein phosphorylation and signaling. Cell. 1995; 80:225-36. [PMID: 7834742]
  45. Lee J, Xu Y, Chen Y, Sprung R, Kim SC, Xie S, Zhao Y. Mitochondrial phosphoproteome revealed by an improved IMAC method and MS/MS/MS. Mol Cell Proteomics. 2007; 6:669-76. [PMID: 17208939]
  46. Jensen SS, Larsen MR. Evaluation of the impact of some experimental procedures on different phosphopeptide enrichment techniques. Rapid Commun Mass Spectrom. 2007; 21:3635-45. [PMID: 17939157]
  47. Thingholm TE, Jensen ON, Robinson PJ, Larsen MR. SIMAC (sequential elution from IMAC), a phosphoproteomics strategy for the rapid separation of monophosphorylated from multiply phosphorylated peptides. Mol Cell Proteomics. 2008; 7:661-71. [PMID: 18039691]
  48. Henzel WJ, Billeci TM, Stults JT, Wong SC, Grimley C, Watanabe C. Identifying proteins from two-dimensional gels by molecular mass searching of peptide fragments in protein sequence databases. Proc Natl Acad Sci USA. 1993; 90:5011-5. [PMID: 8506346]
  49. Ferrini W, Schorderet DF, Othenin-Girard P, Uffer S, Heon E, Munier FL. CRYBA3/A1 gene mutation associated with suture-sparing autosomal dominant congenital nuclear cataract: a novel phenotype. Invest Ophthalmol Vis Sci. 2004; 45:1436-41. [PMID: 15111599]
  50. Cohen D, Bar-Yosef U, Levy J, Gradstein L, Belfair N, Ofir R, Joshua S, Lifshitz T, Carmi R, Birk OS. Homozygous CRYBB1 deletion mutation underlies autosomal recessive congenital cataract. Invest Ophthalmol Vis Sci. 2007; 48:2208-13. [PMID: 17460281]
  51. Zhang LY, Gong B, Tong JP, Fan DS, Chiang SW, Lou D, Lam DS, Yam GH, Pang CP. A novel gammaD-crystallin mutation causes mild changes in protein properties but leads to congenital coralliform cataract. Mol Vis. 2009; 15:1521-9. [PMID: 19668596]
  52. Sabri A, Rafiq K, Seqqat R, Kolpakov MA, Dillon R, Dell'italia LJ. Sympathetic activation causes focal adhesion signaling alteration in early compensated volume overload attributable to isolated mitral regurgitation in the dog. Circ Res. 2008; 102:1127-36. [PMID: 18356543]
  53. Lopes LB, Furnish EJ, Komalavilas P, Flynn CR, Ashby P, Hansen A, Ly DP, Yang GP, Longaker MT, Panitch A, Brophy CM. Cell permeant peptide analogues of the small heat shock protein, HSP20, reduce TGF-beta1-induced CTGF expression in keloid fibroblasts. J Invest Dermatol. 2009; 129:590-8. [PMID: 18787533]
  54. Asomugha CO, Gupta R, Srivastava OP. Identification of crystallin modifications in the human lens cortex and nucleus using laser capture microdissection and CyDye labeling. Mol Vis. 2010; 16:476-94. [PMID: 20352024]
  55. Wang Z, Obidike JE, Schey KL. Posttranslational modifications of the bovine lens beaded filament proteins filensin and CP49. Invest Ophthalmol Vis Sci. 2010; 51:1565-74. [PMID: 19875662]