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

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 BCA^TM 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 BCA^TM 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 ZipTips^TM (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-TOF^TM Premier; Waters Corp, Milford, MA). For LC-MS/MS analysis by Waters Q-TOF^TM 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 LC^TM 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-TOF^TM 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.

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.

Conclusions

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.