Evolution of Src Homology 2 (SH2) Domain to Recognize

Jul 18, 2016 - Specifically, we created a mutant library that was based on the SH2 domain from the proto-oncogene tyrosine-protein kinase Src. To simp...
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Evolution of Src Homology 2 (SH2) Domain to Recognize Sulfotyrosine Tong Ju,† Wei Niu,*,‡ and Jiantao Guo*,† †

Department of Chemistry, University of NebraskaLincoln, Lincoln, Nebraska 68588, United States Department of Chemical & Biomolecular Engineering, University of NebraskaLincoln, Lincoln, Nebraska 68588, United States

‡

S Supporting Information *

ABSTRACT: Protein tyrosine O-sulfation is considered as the most common type of post-translational tyrosine modification in nature and plays important roles in extracellular biomolecular interactions. To facilitate the mapping, biological study, and medicinal application of this type of post-translational modification, we seek to evolve a small protein scaffold that recognizes sulfotyrosine with high affinity. We focused our efforts on the engineering of the Src Homology 2 (SH2) domain, which represents the largest class of known phosphotyrosine-recognition domain in nature and has a highly evolvable binding pocket. By using phage display, we successfully engineered the SH2 domain to recognize sulfotyrosine with high affinity. The best mutant, SH2-60.1, displayed more than 1700 fold higher sulfotyrosine-binding affinity than that of the wild-type SH2 domain. We also demonstrated that the evolved SH2 domain mutants could be used to detect sulfoprotein levels on the cell surface. These evolved SH2 domain mutants can be potentially applied to the study of protein tyrosine O-sulfation with proper experimental designs.

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rotein tyrosine O-sulfation is considered the most common type of post-translational tyrosine modification in nature.1,2 As the only type of direct protein sulfation that is unequivocally confirmed,3,4 it occurs exclusively on secreted and membrane-bound proteins that transit the trans-Golgi network.1,5,6 The introduction of a negatively charged sulfate group plays crucial roles in extracellular biomolecular interactions that dictate various cellular processes including cell adhesion, leukocyte trafficking, hormone activities, and immune responses.2−4,7 It was shown that tyrosylprotein sulfotransferases (TPSTs) knockout mice either died (94%) in the early postnatal period due to cardiopulmonary insufficiency or displayed primary hypothyroidism.8 Tyrosinesulfated proteins (referred to hereafter as sulfoproteins) also play important roles in the development of infectious diseases (e.g., AIDS9,10 and malaria11), cancers, and a variety of immune-mediated diseases (e.g., allergy, rheumatoid arthritis, and inflammatory bowel disease).12 Hence, protein tyrosine O-sulfation could emerge as an important therapeutic target for the treatment of human diseases. However, the lack of understanding on the role and the extent of protein tyrosine O-sulfation in mammalian cell biology severely hinders protein sulfation-associated therapeutic interventions. For example, a search of the UniProt database revealed less than 50 known human sulfoproteins, which account for only 0.08% of the human proteome. This number is much less than the bioinformatics estimate,13,14 suggesting up to 1% of total tyrosine residues have undergone sulfation in animals.1,2 Since sulfotyrosine (sTyr; Figure 1A) is extremely labile under mass spectrometry conditions, it remains © XXXX American Chemical Society

Figure 1. Structures of sulfotyrosine (sTyr), phosphotyrosine (pTyr), and modified peptides. (A) Structures of sTyr and pTyr. (B) Structures of peptides that were used in this study. The biotinylated peptides, biotin-sulfopeptide, biotin-phosphopeptide, and biotintyrosinepeptide, were used in the biopanning. The fluorescein-labeled peptides, FITC-sulfopeptide, FITC-phosphopeptide, and FITCtyrosinepeptide, were used in the fluorescence polarization assays.

Received: June 24, 2016 Accepted: July 18, 2016

A

DOI: 10.1021/acschembio.6b00555 ACS Chem. Biol. XXXX, XXX, XXX−XXX

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ACS Chemical Biology

mutations would improve sTyr binding as well, since both sTyr and pTyr contain an identical phenyl ring. The resulting SH2 domain, SH2-tm, was used as the template for library construction. On the basis of structures23,24 of Src SH2 domain in complex with a phosphopeptide23,24 and results from our previous investigation on SH2 domain-sTyr interaction,19 five residues (Arg35, Glu36, Ser37, Glu38, Thr39) that have potential interaction with the phosphate group of pTyr were chosen for randomization (Figure 2). All five residues reside within the pTyr binding pocket of the Src SH2 domain and do not have apparent interaction with other residues from the target sulfopeptide substrate.

technically challenging to identify sulfoproteins in a crude cell lysate.15 Hence, the scarcity of reports on sulfoproteins can likely be attributed to an analytical problem rather than to a low abundance of this type of post-translational modification. In order to conduct mass spectrometry analysis of sulfoproteins, it requires a high level of enrichment of sulfoproteins from a large excess of unsulfurylated background proteins. Anti-sTyr antibodies have proven to be promising in this regard. Two anti-sTyr antibodies have been generated.16,17 While anti-sTyr antibodies work well in common biological applications (e.g., Western blot), their affinity toward sTyr and the cost are not satisfactory in certain studies (e.g., proteomics) based on our own experiences. Here, we report our recent work on the engineering of Src homology 2 (SH2) domain for the recognition of sTyr. In comparison to anti-sTyr antibodies, the evolved SH2 domain mutants cannot differentiate between sTyr and phosphotyrosine (pTyr), but have advantages in affinity and cost. Under proper experimental conditions (e.g., removing pTyr interference in the sample through phosphatase treatment), these SH2 domain mutants can facilitate the study of protein tyrosine O-sulfation (e.g., sulfoproteome).

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RESULTS AND DISCUSSION There are four major reasons that the SH2 domain was chosen as the target protein scaffold for engineering: (1) The first is the small size (∼12 kDa). In certain applications, the sTyr-binding SH2 domain could potentially be a good alternative choice to the anti-sTyr antibody, which has a low affinity and a large size. (2) Next is similar ligand structures. The SH2 domain naturally recognizes phosphotyrosine (pTyr)-containing protein/peptide. It is known that the pTyr residue makes a major contribution to the binding free energy (ΔG°) of a phosphopeptide to an SH2 domain.18 pTyr and sTyr share a similar structure. Indeed, the Src SH2 domain displayed a low but measurable affinity toward FITC-sulfopeptide (Figure 1B; KD = 9620.0 ± 1670.0 nM).19 (3) Next is the evolvable nature. Previous work has shown that SH2 domains were highly evolvable toward their pTyr-containing substrates.20 We also demonstrated that the affinity of the Src SH2 domain toward sulfopeptide could be slightly improved through point mutations.19 (4) Last is the tunable binding mechanism. By engineering only the pTyr-binding pocket, SH2 domain mutants could be generated with significantly improved affinity toward pTyr in a sequence-independent manner.20 Using a similar engineering approach, protein/peptide sequenceindependent sTyr-binding SH2 domain mutants could potentially be obtained for certain applications, such as sulfoproteome studies. Construction of SH2 Domain Library. A phage displaybased directed evolution approach21 was used in this study to generate and to select SH2 domain mutants with desired properties. Specifically, we created a mutant library that was based on the SH2 domain from the proto-oncogene tyrosineprotein kinase Src. To simplify the phage display process, we mutated two cysteine residues (Cys98 and Cys105) of the wildtype Src SH2 domain (SH2-wt) into serine residues. According to literature report,22 such mutations do not compromise the structure and substrate-binding properties of the SH2 domain. In addition, we introduced three previously discovered mutations (Thr40Val, Cys45Ala, and Lys63Leu),20 which form a hydrophobic surface to beneficially engage the phenyl ring of the pTyr residue and render Src SH2 domain mutants with higher affinity toward pTyr. We envisaged that these three

Figure 2. Crystal structure of the Src SH2 domain in complex with a phosphopeptide (PDB: 1SPS). The phosphopeptide is shown in yellow.

Biopanning. The SH2 domain library was displayed on the surface of the M13 bacteriophage and screened for the ability to bind to a sulfopeptide (biotin-sulfopeptide; Figure 1B) substrate, which was immobilized in the wells of a streptavidin coated 96-well plate. This biotin-sulfopeptide has the amino acid sequence of the natural substrate of the Src SH2 domain but contains sTyr at the phosphorylation site. Two different selection schemes were employed. In the first one, five rounds of consecutive positive selections against biotin-sulfopeptide were conducted. In the second selection scheme, alternate positive selection against biotin-sulfopeptide and negative selection against biotin-phosphopeptide (Figure 1B) were performed. Negative selections against unsulfated peptides were not included in our two selection schemes since we did not obtain mutants with improved binding affinity when the library was selected toward the biotin-tyrosinepeptide (Figure 1B). We hypothesized that the negatively charged sulfate or phosphate group was the key recognition element for our SH2 domain library. Following five to six rounds of selection, phage enrichment reached a plateau. Data of phage ELISA experiments showed that the enriched phage library displayed significantly higher affinity toward the target biotin-sulfopeptide than that of the naive phage library. Single phage clones were then isolated and characterized in semiquantitative phage ELISA experiments. DNA sequences of eight clones with the highest affinity were analyzed (Table 1). Within these eight candidates, six unique sequences were obtained while three clones, SH2-60.1, SH269.1, and SH2-69.4, converged to the same sequence (Table 1). Further characterizations were conducted with the top two hits, SH2-60.1 (from alternate positive/negative selection scheme) B

DOI: 10.1021/acschembio.6b00555 ACS Chem. Biol. XXXX, XXX, XXX−XXX

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the SH2-wt and SH2-tm, respectively. As a second comparison, under the same assay conditions, the KD value of a commercially available anti-sTyr antibody (Clone: Sulfo-1CA2, EMD Millipore) toward FITC-sulfopeptide was apparently higher than 1000 nM (Figure S3). As a third comparison, the KD value of SH2-wt toward its native FITC-phosphopeptide substrate is about 61 nM.19 None of the SH2 domain variants, including SH2-wt, SH2tm, SH2-60.1, and SH2-58.6, displayed measurable affinity toward FITC-tyrosinepeptide. This result demonstrates that the evolved SH2 mutants have excellent selectivity toward sTyr over Tyr. It is clear that sTyr is the major recognition element by the evolved SH2 domain mutants, while other residues on the substrate peptide do not contribute significantly to the binding free energy. We therefore hypothesized that the evolved SH2 mutants should bind to sulfoproteins in a relatively sequence-independent manner, a trait that is necessary for sulfoproteome studies. On the other hand, none of the SH2 domain variants (SH2-tm, SH2-60.1, or SH2-58.6) was able to differentiate between sTyr and pTyr (Figure 3). It was to our surprise that SH2-60.1 recognized sTyr and pTyr with similar affinity, although it was obtained from a selection scheme in which three rounds of negative selections against biotin-phosphopeptide were included. A number of negative selection conditions were subsequently examined. But SH2 domain mutants that could efficiently differentiate sTyr from pTyr were not obtained. Ongoing experiments are conducted to further investigate this problem. In the mean time, whole-cell sulfoproteome studies can perceivably be performed by applying the evolved SH2 domain mutants to protein tyrosine phosphatase-treated total cellular protein fraction.25 We will evaluate this application in the near future. Besides proteins, carbohydrate moieties on proteins and lipids are also subjected to sulfation. A previous study showed

Table 1. Sequence Analysis of the SH2 Domain Variants SH2 variants SH2-tm SH2-58.2 SH2-58.5 SH2-58.6 SH2-60.1 SH2-69.1 SH2-69.4 SH2-69.5 SH2-6.8.1

sequence Arg35 Pro Gly Arg Arg Arg Arg Arg Arg

Glu36 Trp Asp Gln Arg Arg Arg Gln Arg

Ser37 Trp Gln Leu Leu Leu Leu Val Gln

Glu38 Asn Gln Ala Gln Gln Gln Arg Arg

Thr39 Gln His Arg Arg Arg Arg Lys Pro

and SH2-58.6 (from the consecutive positive selection scheme). Characterization and Validation of SH2 Domain Variants. SH2 domain variants, including SH2-wt, SH2-tm, SH2-60.1, and SH2-58.6, were partially purified by affinity chromatography (Figure S1). Fluorescence polarization (FP) experiments were conducted to directly measure their KD values toward three peptide substrates, FITC-sulfopeptide, FITC-phosphopeptide, and FITC-tyrosinepeptide. As we expected, the affinity of SH2-tm toward FITC-sulfopeptide was significantly higher than that of SH2-wt (Figure 3). The three beneficial mutations (Thr40Val, Cys45Ala, and Lys63Leu) for the binding of pTyr indeed also improved SH2 domain’s affinity toward sTyr, possibly due to hydrophobic interactions.20 The two evolved SH2 domain mutants, SH260.1 and SH2-58.6, showed excellent affinity (KD = 5.6 ± 0.8 and 10.4 ± 1.0 nM; Figure 3) toward FITC-sulfopeptide. As a comparison, the KD values of SH2-wt and SH2-tm toward FITC-sulfopeptide are 9620.0 ± 1670.0 nM and 348.5 ± 64.5 nM, respectively. Therefore, the best mutant, SH2-60.1, displayed more than 1700-fold and 60-fold improvement over

Figure 3. Recognition of sulfopeptides by the Src SH2 domain variants. The KD values were obtained by fluorescence polarization assays. Each data point is the average of triplicate measurements with standard deviation. C

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ACS Chemical Biology that the wild-type Src SH2 domain could similarly recognize both the sulfogalactose on sulfoglycosphingolipids and sTyr.26 Therefore, sTyr and sulfogalactose were hypothesized to be mutual mimicry in that study.26 Although our FP data indicated a very weak interaction (Kd = 9620 nM; Figure 3) between the Src SH2 domain and sulfopeptide, due to the natural abundance of sulfated sugar, in particular on the cell surface, the binding affinity of SH2 domain variants toward three sulfated sugars was investigated. Competitive fluorescence polarization experiments were conducted by including varied concentrations of sulfate sugars in regular FP assays. As shown in Figure S6, no apparent interference to the SH2 domain (0.2 μM)/FITC-sulfopeptide (0.01 μM) interaction was observed in the presence of high concentrations of sulfated carbohydrates (0.48−1000 μM). On the other hand, the binding between the SH2 domain and FITC-sulfopeptide was nearly completely disrupted in the presence of 0.05 μM of biotin-sulfopeptide (Figure S6). All evolved SH2 domain mutants contain more positively charged residues than that of SH2-tm (Table 1). With additional positively charged residues, the evolved SH2 domain mutants are likely able to form stronger electrostatic interactions with the negatively charged sTyr and, therefore, displayed higher affinity toward sTyr than that of SH2-wt and SH2-tm. On the other hand, the evolved SH2 domain mutants and their parent, SH2-tm, showed similar affinity toward pTyr (Table 1). The beneficial effects of additional positively charged residues are either not necessary for pTyr binding or offset by topological distortion of the optimal binding pocket toward pTyr. Detection of Cell Surface Sulfoprotein Levels. Different cell types contain characteristic sets of sulfoproteins. Protein sulfation level of a cell could also vary under different growth/ pathological conditions. In addition, the change of sulfation level of cell surface receptors affects many downstream signaling pathways.9,27−29 We therefore examined the evolved SH2 domain variants for their utility in rapid assessment of cell surface sulfoprotein level. To test if the evolved SH2 domain mutants can be used to detect sulfoproteins on the cell surface, three SH2 domain variants, including SH2-wt, SH2-60.1, and SH2-58.6, were fused to the C-terminus of green florescent protein (GFP). The three GFP-SH2 fusion proteins (GFP-SH2-wt, GFP-SH2-60.1, and GFP-SH2-58.6) were expressed in E. coli and purified by affinity chromatography (Figure S2). The purified fusion proteins were incubated with 293T cells for 1.5 h. After removal of the culture medium, cells were washed with DPBS buffer, fixed with 4% paraformaldehyde (w/v) solution, and visualized under a confocal microscope. As shown in Figure 4, intense fluorescence was observed on the cell surface of samples that were treated with GFP-SH2-60.1 or GFP-SH2-58.6. As a control, no fluorescence labeling was detected when GFP-SH2wt fusion protein was used in the labeling experiment. Since the only difference between the SH2 domain mutants and the wildtype SH2 domain is the evolved binding pocket, the results indicated that the observed fluorescence labeling was likely due to the binding of the evolved SH2 domain mutants to sTyr residues of sulfoproteins. Nonspecific bindings to sulfated sugars on cell surfaces were not supported by data of our competitive FP assays. We further demonstrated that cell surface labeling by SH2 domain mutants was not affected in the presence of a high concentration (10 000 μM) of sulfated carbohydrates (Figures S7, S8). It is also unlikely that the

Figure 4. Labeling of cell-surface sulfoproteins with GFP-SH2 fusion protein variants. The first column shows fluorescence images. The second column shows bright-field images. The third column shows composite images. The protein concentration was 5 μM for all three fusion proteins.

labeling signal was a result of bindings of SH2 domain mutants to pTyr-containing proteins. To the best of our knowledge, only two potential pTyr-containing proteins were reported in a special cell line (K562 human leukemic cell).30 The notion of a lower level of pTyr-containing proteins on the cell surface was supported by our cell-based ELISA experiments using both anti-sTyr and anti-pTyr antibodies, in which a more than 7-fold lower signal was obtained with anti-pTyr antibodies (Figure S5). Overall, the above data suggested that the observed signal in Figure 4 likely resulted from the detection of sTyr-containing proteins, and not from pTyr-containing proteins or sulfated carbohydrates. To demonstrate that the evolved SH2 domains can be used to analyze sulfoprotein levels on the cell surface, cells with various levels of sulfoproteins were obtained by the treatment with different concentrations of sodium chlorate (NaClO3).31−34 As an analog of sulfate, chlorate inhibits ATP-sulfurylase, which is an essential enzyme in the biosynthesis of the universal sulfate donor, 3′-phosphoadenosine 5′phosphosulfate (PAPS). Up to 50 mM NaClO3 has been used to effectively reduce the sulfation of both proteins and proteoglycans.31−34 Protein sulfation could be nearly completely inhibited when higher than 20 mM NaClO3 was used.34 In a control experiment with an anti-sTyr antibody, we confirmed the inverse correlation between sulfation levels of cell surface proteins and the concentrations of NaClO3 that were used to treat cells (Figure S4). Next, we examined the ability of evolved SH2 domains to detect protein sulfation level. Following the treatment with 0, 10, 30, and 50 mM NaClO3, cells were washed and incubated with GFP-SH2-60.1 or GFPSH2-58.6 for 1.5 h in fresh culture media. The cells were then washed and visualized under a confocal microscope. As shown in Figure 5, significantly lower fluorescence intensities were observed with cells treated by NaClO 3 . In addition, fluorescence signal decreased as the NaClO3 concentration increased (Figure 5). A very low level of fluorescence was detected in samples that were treated with 30 mM and 50 mM of NaClO3, presumably because of the extremely low levels of sulfoproteins as reported in the literature.34 These results further supported our conclusion that the labeling signals by GFP-SH2 fusion proteins were not caused by binding of SH2 domain mutants to pTyr on phosphoproteins, of which the expression level is not affected by chlorate. Consistently D

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Figure 5. Labeling of cell-surface sulfoproteins with GFP-SH2 variants after the treatment of cells with different concentrations of NaClO3. In each panel, the first column shows fluorescence images, the second column shows bright-field images, and the third column shows composite images. The protein concentration was 5 μM for both fusion proteins. SH2 Domain Library Construction. Procedures for library construction were adapted from Noren and Noren.36 Template DNA of SH2-tm was generated by site-directed mutagenesis to install C98S/C105S and T40V/C45A/K63L mutations.20 DNA sequences encoding selected residues (Arg35, Glu36, Ser37, Glu38, Thr39) were diversified using degenerate primers in overlapping PCR with SH2-tm as the template. Digestion of PCR products with EcoRI and BamHI followed by ligation into pFN-OM6 vector treated with the same restriction enzymes resulted in the SH2 domain library, pFN-OM6-lib. Sequences of primers are reported in the Supporting Information. Phage Display and Biopanning. To generate phages that display the SH2 domain library, E. coli XL1-Blue cells were transformed with pFN-OM6-lib and infected by M13KO7 helper phage. Biopanning for positive selection was performed by incubation of the phage library (1012 phage/mL) with biotin-sulfopeptide (Figure 1B) immobilized in a well of a streptavidin coated 96-well plate (Thermo Scientific). After 1 h of incubation, the well was washed 10 times using PT buffer (PBS with 0.05% Tween); the bound phages were eluted and amplified by infecting XL1-Blue cells. A small portion of the infected XL1-Blue cells was used for phage tittering on LB agar plates containing 100 μg/mL ampicillin. The remaining infected cells were subsequently cultivated at 37 °C overnight in 2 × YT medium containing 100 μg/mL ampicillin, 50 μg/mL kanamycin, and 0.1 mM IPTG. The cells were then pelleted down, and the phage pool was precipitated from the supernatant using 20% PEG/NaCl solution. The amplified phage pool was subsequently used for the next round of selection. Biopanning for negative selection was performed by incubation of the phage library (1012 phage/mL) with biotin-phosphopeptide (Figure 1B) immobilized in a well of a streptavidin coated 96-well plate. After 1 h of incubation, the unbound phages were collected in PT buffer and amplified by infecting XL1-Blue cells. A small portion of the infected XL1-Blue cells was used for phage tittering on LB agar plates containing 100 μg/mL ampicillin. The remaining infected cells were subsequently cultivated at 37 °C overnight in 2 × YT medium containing 100 μg/mL ampicillin, 50 μg/mL kanamycin, and 0.1 mM IPTG. The cells were then pelleted down, and the phage pool was precipitated from the supernatant using 20% PEG/NaCl solution. The amplified phage pool was then used for the next round of selection. Two different selection schemes were employed. In the first one, five rounds of consecutive positive selections against biotinsulfopeptide were conducted. In the second selection scheme, alternate positive selection against biotin-sulfopeptide and negative selection against biotin-phosphopeptide were performed. The biopanning process was repeated five to six times to enrich phages that have high affinity toward biotin-sulfopeptide. Individual phage clones were

stronger signals were observed when GFP-SH2-60.1 (relative to SH2-58.6) was applied as the detecting agent. This result agrees with the fact that SH2-60.1 has a higher affinity toward sulfoproteins than that of SH2-58.6. The above data demonstrated that GFP-SH2 fusion proteins could provide direct visual information on the sulfoprotein level through a simple protocol. Other reporter proteins, such as luciferase and HRP, can perceivably be fused to the evolved SH2 domain in order to fit the experimental needs. Conclusion. In summary, we demonstrated that the SH2 domain could be evolved to recognize sTyr with excellent affinity in a sequence-independent manner. The evolved SH2 domain mutants could be used to detect the sulfoprotein level on the cell surface. Efforts are being devoted to the engineering of SH2 domain mutants that can differentiate sTyr and pTyr, which will greatly expand their utility and simplify experimental designs in studies such as sulfoproteome. The evolution of sequence-specific SH2 domain mutants, which can be used to study and regulate individual sulfoproteins with important biological function, is another focus of our ongoing efforts. The high affinity, small size, and easy accessibility of the evolved SH2 domain mutants make them potential alternatives to the anti-sTyr antibody in studies of protein tyrosine O-sulfation.

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METHODS

Materials and General Methods. Peptides were purchased from NeoBioLab, Inc. The pFN-OM6 vector is a gift from S. S. Sidhu at the University of Toronto. Primers were ordered from Sigma. The M13KO7 helper phage, restriction enzymes, Antarctic phosphatase (AP), and T4 DNA ligase were purchased from New England Biolabs. KOD hot start DNA polymerase was purchased from EMD Millipore. Standard molecular biology techniques35 were used throughout. The HRP/anti-M13 monoclonal antibody conjugate was obtained from GE Health Care Life Science. Antisulfotyrosine antibody was purchased from EMD Millipore. Site-directed mutagenesis was carried out using overlapping PCR. E. coli GeneHogs were used for routine cloning and DNA propagation. E. coli XL1-Blue cells were used for phage propagation. E. coli BL21(DE3) cells were used for protein expression. All solutions were prepared in deionized water further treated by a Barnstead Nanopure ultrapure water purification system. Antibiotics were added where appropriate to the following final concentrations: ampicillin, 100 mg L−1; kanamycin, 50 mg L−1; tetracycline, 12.5 mg L−1. E

DOI: 10.1021/acschembio.6b00555 ACS Chem. Biol. XXXX, XXX, XXX−XXX

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Analyses of Sulfoprotein Levels. Sulfoprotein levels were reduced by inhibiting PAPS biosynthesis using NaClO3.32,33 To do this, 293T cells, which were grown in media containing DMEM, 10% FBS (v/v), and 2 mM L-glutamine at 37 °C in a humidified atmosphere of 5% CO2 (v/v), were treated with 0, 10, 30, or 50 mM of NaClO3 in DMEM with 10% FBS at 37 °C for 36 h. When confluence was achieved, the pretreated cells were transferred to a 24well plate and cultivated in the presence of 0, 10, 30, or 50 mM NaClO3 for an additional 12 to 15 h. When cells reached 60−70% confluency, cell culture media were changed to fresh DMEM media (with 10% FBS and 2 mM L-glutamine) containing 5 μM of a GFPSH2 domain fusion protein of interest. After incubation for 1.5 h at 37 °C, the culture medium was removed and replaced with fresh DMEM medium. The cells were incubated for 10 min at 37 °C, then washed with DPBS once. After fixation with 4% paraformaldehyde (w/v), cells were visualized by an inverted confocal microscope (Olympus FV500 System on IX81 scope). In the cases of competitive cell labeling experiments, cells were incubated with GFP-SH2 domain fusion proteins in the presence of 10 000 μM of sulfated carbohydrates. Further processing of the samples followed the same procedure as described above.

then separated, and the interaction between the phage-displayed SH2 domain variants and sulfopeptide were further evaluated by phage enzyme-linked immunosorbent assay (phage ELISA). A few positive clones were subject to DNA sequencing analysis to determine the mutations of the evolved SH2 domain variants. Phage Enzyme-Linked Immunosorbent Assay (Phage ELISA). Individual phages in 100 μL of PBT buffer (PT buffer with 0.5% BSA; 0.8 × 1012 phage/mL) were incubated with biotinsulfopeptide immobilized in the streptavidin-coated wells at RT for 1 h. After the wells were washed eight times with PT buffer (200 μL each time), HRP/anti-M13 monoclonal antibody conjugate (100 μL, 1:5000 dilution) in PBT buffer was added to each well, and the resulting mixtures were incubated for 1 h. The wells were washed six times with PT buffer and two times with PBS buffer, and the 1-Step Slow TMB-ELISA Substrate (100 μL, Thermo Scientific) was added to each well. After shaking at RT for 15 min, the reaction was stopped by the addition of 2 M sulfuric acid (50 μL), and the absorbance at 450 nm was measured. Protein Expression and Purification. SH2 domain variants were subcloned into NdeI and XhoI sites of pET30b to afford plasmid pET30b-SH2-wt, pET30b-SH2-tm, pET30b-SH2-60.1, and pET30bSH2-58.6. These SH2 domains of interest were expressed as fusion proteins containing a C-terminal 6 × His tag. Genes that encode GFPSH2 domain fusion proteins were constructed through overlapping PCR. The fusion protein encoding genes were inserted between the NdeI and XhoI sites of pET30b to afford plasmid pET30b-GFP-SH2wt, pET30b-GFP-SH2-60.1, and pET30b-GFP-SH2-58.6. In these constructs, SH2 domains were fused to the C-terminus of GFP. E. coli BL21(DE3) cells harboring either pET30b-SH2 or pET30bGFP-SH2 variants were cultured in 500 mL of LB medium containing 50 μg/mL kanamycin at 37 °C with agitation. When OD600 reached 0.4−0.6, protein expression was induced by the addition of IPTG to a final concentration of 0.2 mM. After an additional 18 h of growth at 37 °C (or at RT), cells were harvested and lysed by sonication. Cellular debris was removed by centrifugation (21 000g, 30 min, 4 °C). The cell-free lysate was applied to Ni Sepharose 6 Fast Flow resin (GE Healthcare). Protein purification followed the manufacturer’s instructions. Purified protein was desalted using the Econo-Pac 10DG desalting column (BioRad). Protein concentrations were determined by Bradford assay (Bio-Rad). Fluorescence Polarization Assay. Three fluorescein-labeled peptide probes, FITC-sulfopeptide, FITC-phosphopeptide, and FITC-tyrosinepeptide (FITC, fluorescein isothiocyanate), were used. Individual peptide probes were dissolved in assay buffer containing potassium phosphate (20 mM, pH 7.35), NaCl (100 mM), DTT (2 mM), and bovine gamma globulin (0.1%). Aliquots of the peptide probe solution were distributed to 96-well fluorescence plate to reach a final concentration of 10 nM. After the addition of indicated amounts of SH2 domain protein, the assay solutions were mixed and incubated at RT in the dark for 25 min. Fluorescence polarization experiments were performed on a Synergy H1 plate reader (BioTek Instruments, Inc.) equipped with standard filter cube (λEx = 485 nm, BP = 20 nm; λEm = 528 nm, BP = 20 nm). A standard sample layout in a 96-well plate was designed so that a single plate contains all the samples for one SH2 domain variant. Triplicate samples at each protein concentration were arrayed in the plate, together with control samples containing either only the SH2 domain or only the peptide probe. Fluorescence intensity data were measured, exported, and converted into anisotropy values. Calculation of the percentage of bound probe followed reported method.37 Dissociation constants were calculated by curve fitting into a one-site-specific binding equation with a Hill slope using GraphPad Prism 5. Fluorescence polarization assays of all SH2 domain variants were repeated three times. Results were reported as the average with standard deviation. In the cases of competitive fluorescence polarization assays, SH2 domain variants (0.2 μM) were incubated with serial dilutions of competitors (for sulfated carbohydrates, 0.48−1000 μM; for biotin-sulfopeptide, 0.01−1.0 μM) and 0.01 μM of FITC-sulfopeptide probe in assay buffer. After incubation at RT for 15 min, samples were examined using a plate reader as described above.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acschembio.6b00555. Primer list and additional data (PDF)

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This research was supported by the New Faculty Startup Fund (to J.G. and W.N.) and an Interdisciplinary Grant (to J.G.) from the University of NebraskaLincoln. The authors thank Y. Zhou in the Microscopy facility at the Center for Biotechnology in University of NebraskaLincoln for help in fluorescence microscope analysis. We also thank S. S. Sidhu (University of Toronto) for providing plasmid pFN-OM6.

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DOI: 10.1021/acschembio.6b00555 ACS Chem. Biol. XXXX, XXX, XXX−XXX