Accepted Manuscript Expression, purification, and characterization of human osteoclastic proteintyrosine phosphatase catalytic domain in Escherichia coli Huan Jiang, Sui Yuan, Cui Yue, Peng Lin, Wannan Li, Shu Xing, Deli Wang, Min Hu, Xueqi Fu PII: DOI: Reference:
S1046-5928(14)00269-1 http://dx.doi.org/10.1016/j.pep.2014.11.008 YPREP 4604
To appear in:
Protein Expression and Purification
Received Date: Revised Date:
3 September 2014 10 November 2014
Please cite this article as: H. Jiang, S. Yuan, C. Yue, P. Lin, W. Li, S. Xing, D. Wang, M. Hu, X. Fu, Expression, purification, and characterization of human osteoclastic protein-tyrosine phosphatase catalytic domain in Escherichia coli, Protein Expression and Purification (2014), doi: http://dx.doi.org/10.1016/j.pep.2014.11.008
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Expression, purification, and characterization of human osteoclastic protein-tyrosine phosphatase catalytic domain in Escherichia coli Huan Jianga, Sui Yuanb, Cui Yuea, Peng Lina,Wannan Lib, Shu Xingb, Deli Wangc, Min Hua*, Xueqi Fub*
a
Department of Orthodontics, School of Stomatology, Jilin University, 1500 Qinghua Road, Changchun 130021, P. R. China. b
Edmond H. Fischer Signal Transduction Laboratory, School of Life Sciences, Jilin University, Changchun 130012, P. R. China. c
School of Life Sciences, Jilin University, Changchun 130012, P. R. China.
The authors report no commercial, proprietary, or financial interest in the products or companies described in this article. Supported by the National Natural Science Foundation of China (811709999/H1409).
*Corresponding author: Min Hu, Department of Orthodontics, College of Stomatology, Jilin University. No. 1500, Qinghua Rd, Changchun 130021, Jilin, P. R. China; E-mail: [email protected]. Tel.: +86-431-887-96023; Fax: +86-431-889-75348.
Xueqi Fu, Edmond H. Fischer Signal Transduction Laboratory, College of Life Sciences, Jilin University, No. 2699, Qianjin Street,Changchun 130021, Jilin, P. R. China; E-Mail: [email protected]. Tel.: +86-431-851-55321; Fax: +86-431-851-55152.
ABSTRACT Osteoclastic protein tyrosine phosphatase (PTP-oc) is a structurally unique transmembrane protein tyrosine phosphatase (PTP) that contains only a relatively small intracellular PTP catalytic domain, does not have an extracellular domain, and lacks a signal peptide proximal to the NH2 terminus. The present study reports the expression, purification, and characterization of the intracellular catalytic domain of PTP-oc (∆PTP-oc). ∆PTP-oc was expressed in Escherichia coli cells as a fusion with a six-histidine tag and was purified via nickel affinity chromatography. When using para-nitrophenylphosphate (p-NPP) as a substrate, ∆PTP-oc exhibited classical Michaelis-Menten kinetics. Its responses to temperature and ionic strength were similar to those of other PTPs. The optimal pH value of ∆PTP-oc is approximately 7.0, unlike other PTPs, whose optimal pH values are approximately 5.0.
Keywords: Osteoclastic protein tyrosine phosphatase, Escherichia coli, Nickel affinity chromatography, Enzyme kinetics
Introduction Intracellular tyrosyl-phosphorylation levels are tightly regulated by protein tyrosine kinases (PTKs) and protein tyrosine phosphotases (PTPs) [1]. In this process, the phosphorylation reaction is mediated by PTKs, whereas the dephosphorylation reaction is catalyzed by PTPs [1,2]. The importance of tyrosine phosphorylation is illustrated best in osteoclasts [6,7], when considering the major signaling pathways, such as those mediated by RANKL and M-CSF [8,9]. Molecular studies indicate that phosphorylation of proteins on tyrosine residues is critical for the production and function of osteoclasts [2,3,4,5]. However, beyond these roles, compelling evidence also supports protein tyrosine phosphorylation having essential regulatory functions in integrin signaling in osteoclasts [10-12]. Osteoclastic protein tyrosine phosphatase (PTP-oc) belongs to the receptor PTP family and is mainly expressed in precursors of osteoclasts, including B lymphocytes, cells of the monocyte-macrophage lineage, and mature osteoclasts; it is also an activator of osteoclast differentiation and activity [13-16,20,21]. PTP-oc is the product of a gene assigned to human chromosome 12p12-p13 [17,18] or mouse chromosome 6 [19], and the expression of PTP-oc is driven by an intronic, tissue-specific proximal promoter of the PTPRO gene [13,14]. Unlike conventional transmembrane PTPs, PTP-oc is constitutive and consists of only 405 amino acid residues, which is relatively small for a transmembrane protein. The structure of PTP-oc is unique in that it contains only a single intracellular PTP catalytic domain, has a very short extracellular domain and lacks a signal peptide proximal to the NH2 terminus [13,16]. PTP-oc is an interesting protein tyrosine phosphatase, as studies show that it not only displays tissue-specific expression but also has tissue-specific functions [14]. PTP-oc may function as a positive regulator of osteoclast activity, proliferation, and differentiation [16,21 ]. Upon targeted overexpression of wildtype PTP-oc, osteoclast-like cells derived from human monocytic U937 cells created resorption pits that were larger and deeper than those of osteoclast-like cells derived from cells overexpressing phosphatase-deficient PTP-oc [16]. More importantly, more significant trabecular bone loss occurred because of increased bone resorption, but not decreased bone formation, in the osteoclastic lineage overexpressing PTP-oc in vivo [15,16, 20]. PTP-oc
stimulates osteoclasts in part by activating c-Src through dephosphorylation of its Tyr-527 (pY527) residue [15,16, 22-24]. The suppression of PTP-oc expression by a PTP-oc antisense oligodeoxynucleotide in osteoclasts reduced their bone resorption activity and increased their c-Src pY527 level [15,24]. Collectively, these results suggest that PTP-oc dephosphorylates and activates c-Src in osteoclasts; therefore, the downstream effects of this signaling may be mediated via activation of NFêB and JNK2 [22]. In addition to its regulation of osteoclast activity, there is evidence that PTP-oc may regulate osteoclast differentiation by RANKL in vitro [1,25]. Therefore, we highlight the essential regulatory role that PTP-oc may play in osteoclast function. To further study the molecular mechanism of PTP-oc signal transduction, we established an expression and purification system that is capable of providing a large-scale quantity of the functional intracellular catalytic domain of PTP-oc (∆PTP-oc). We cloned the cDNA of ∆PTP-oc into the pET-28a (+) vector, purified the His6-tagged recombinant proteins using nickel affinity chromatography, and performed detailed characterization.
Materials and Methods Construction of expression vector The pET-28a (+) plasmid was used to clone the ∆PTP-oc gene fragment. The cDNA fragment encoding the ∆PTP-oc (nucleotides 870–1656) was amplified by polymerase chain reaction (95°C for 3 min; 94°C for 1 min; 54°C for 1 min; 72°C for 1 min; 72°C for 10 min; 30 cycles) from a human cDNA library (NCBI gene ID: NM_030669.2) and was ligated into the pET-28a (+) vector by T4-DNA ligase (New England Biolabs). The primers were designed to create NcoI and XhoI restriction sites to facilitate the insertion of the ∆PTP-oc fragment into the pET-28a (+) vector. The forward primer was 5′- AGGCGCCATGGGCAAATTTTCTCTTCAGTTTGAGG -3′, and the reverse primer was 5′- GCGATCCTCGAGCACACACTGATGGATAAAAATG -3′. The ligated samples contained a C-terminal nucleotide sequence coding for six histidines, which was placed downstream and in frame with the ∆PTP-oc insert. The sizes of the polymerase chain reaction (PCR) products were confirmed by electrophoresis in a 2% agarose gel. A single colony obtained on Luria-Bertani (LB)-kanamycin plates was selected and cultured. The palsmids were purified and the sequence was confirmed by DNA sequencing. The recombinant plasmids were transformed into BL21 (DE3) cells for protein expression. Expression of His6-tagged ∆PTP-oc BL21 (DE3) cells that harbored the plasmid containing the ∆PTP-oc insert were grown in 2 ml of LB medium supplemented with 50 µg/ml kanamycin. After overnight growth at 37°C with shaking, 2 ml of this pre-culture was used to inoculate 200 ml of fresh LB media containing an appropriate antibiotic in a flask, and the cells were grown at 37°C to mid-log phase. Isopropyl-β-D- thiogalactopyranoside (IPTG) was added at a final concentration of 0.1 mM to induce ∆PTP-oc expression, and the temperature was decreased to 16°C to improve the soluble expression level [30]. After 24 h of growth at 16°C, the cells were harvested at 4°C by centrifugation at 5000 g. The pellets were resuspended in 10 ml/g extraction buffer (50 mM Tris-HCl, pH 7.4, 50 mM NaCl, 10 mM imidazole, 10% glycerol, 20 mM 2-mercaptoethanol, 2% Tween-20) containing a protease inhibitor mixture (1 mM benzamidine, 0.1 mM phenylmethylsulfonyl fluoride, 1 mM pepstatin A, 2 µg/ml leupeptin and 1 µg/ml aprotinin) and broken up by sonication on ice. The lysate was cleared by centrifugation at 13,000 g at 4°C for 30
min and both supernatant and pellets were collected. The expression levels and solubility of ∆PTP-oc were assessed by 12% sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE).
Purification of ∆ PTPPTP -oc The cytosolic supernatant from the above-mentioned step was loaded onto a Ni-NTA agarose column equilibrated with buffer A (50 mM Tris-HCl, pH 7.4, 50 mM NaCl, 10 mM imidazole, 10% glycerol). The column was washed with buffer B (50 mM Tris-HCl, pH 7.4, 500 mM NaCl, 10 mM imidazole, 10% glycerol). The tightly bound proteins were eluted with a linear imidazole gradient of 0–0.2 mol/L, and the protein peak was collected in fractions. The purity of soluble His6-∆PTP-oc was observed on 12% SDS gels. All purification procedures were carried out at 4°C and PTP activity was analyzed by use of para-nitrophenylphosphate (p-NPP) as a substrate.
Protein concentration measurements and SDSSDS-PAGE The protein concentrations were determined using the Bradford method with bovine serum albumin (Sigma) as a reference standard according to the manufacturer’s instructions [31]. Soluble His6-∆PTP-oc was concentrated to 1 mg/ml and separated on 12% Tris-glycine SDS gels. Electrophoresis was performed in a Gibco-BRL Vertical Cell Electrophoresis Apparatus (Life Technologies) with standard molecular mass markers (Transgen Biotech). The gels were stained with Coomassie Blue for 1 h at room temperature. The destaining was performed in 10% acetic acid.
Western blot assays The proteins were separated by 12% SDS-PAGE and then transferred to polyvinylidene difluoride (PVDF) membranes. Western blot analysis was performed by use of an anti-His6 tag monoclonal antibody (Abcam) as primary antibody. Proteins were detected using the enhanced chemiluminescence (ECL) method.
Enzymatic Characterization of ∆ PTPPTP -oc The phosphatase activity of the His6-∆ PTP-oc fusion protein was tested in vitro by p-NPP (Sigma) hydrolysis assays at neutral pH, as previously described [13,27-29]. The reaction mixture consisted of 25 mM MOPS-NaOH (pH 7.0), 1 mM EDTA, 1 mM dithiothreitol, 0.1 mg/ml BSA, 0.05µg/µl purified proteins and p-NPP at the indicated concentrations in a final volume of 100 µl. All activity assays were tested in triplicate. The blanks were prepared in the same manner as the samples but without the enzyme. The reaction was carried out at 37°C for 15 min and terminated by the addition of 100 µl 0.2 M NaOH solution, and the assay results were read against a blank at 405 nm. One unit of activity (U) was defined as 1 nmol of phosphate released per min, and the specific activity was expressed as U/mg [32]. Kinetics of ∆PTP-oc To study the enzyme kinetics of ∆PTP-oc, we performed enzyme assays with various concentrations of p-NPP substrate (S) for 15 min at 37 °C and pH 7.0. The data were processed with Graphpad Prism 4.0.
Results and discussion
Construction of an expression plasmid encoding ∆ PTPPTP-oc with a His tag Primary structure analyses suggest that PTP-oc is a transmembrane protein [13,14,16]. The expression of the full-length enzyme resulted in the distribution of most of the protein in inclusion bodies with relatively little PTP activity in Escherichia coli [32]. This result made the purification of active PTP-oc from E. coli cells unfeasible. The strategy to clone and express recombinant ∆PTP-oc involved inserting the gene of interest into the pET-28a (+) expression vector. Specifically, the insert encoding the phosphatase domain of PTP-oc, which was successfully amplified from the full-length human PTP-oc cDNA from pMD18-T, with the NcoI and XhoI restriction sites, was cloned into the pET-28a (+) vector containing a C-terminal His6 tag to generate pET28a-∆PTP-oc (Fig. 1A). Further characterization using specific restriction endonuclease digests and DNA sequencing demonstrated that the amplified target gene was inserted into the correct open reading frame of the vector (Fig. 1B). The resultant expression construct produced a fusion protein comprising ∆PTP-oc and a LEHHHHHH tag at the C-terminus with a predicted molecular weight of 31.2 kDa, for protein purification by metal affinity chromatography. The ligation products were then transformed into BL21(DE3) competent cells.
Fig. 1. Cloning of pET28a-△PTP-oc. (A) Cloning of pET28a-△PTP-oc from a human cDNA library (NCBI gene ID: NM_030669.2). Lane 1, Molecular weight marker DL-6000; Lane 2, △PTP-oc cloning product. (B) The double digestion of pET28a-△PTP-oc by NcoI and XhoI. Lane 1, Molecular weight marker DL-15000; Lane 2, pET28a-△PTP-oc plasmid; Lane 3, pET28a-△PTP-oc cut by NcoI and XhoI.
Expression of ∆ PTPPTP -oc in E. E . coli The relationship between the temperature dependence of protein induction and inclusion body formation has been extensively studied in a variety of model systems [33,34]. Indeed, an apparent correlation has been noted. Whereby a decrease in the induction temperature gave rise to a lower propensity for recombinant protein expression in inclusion bodies and increased the solubility of recombinant PTPs [33,35,36]. Different temperatures were attempted after induction: 37°C, 32°C, 26°C, 20°C and 16°C. The temperature was finally lowered to 16°C post-induction which highly increased the soluble expression level. After determining the optimal conditions for ∆PTP-oc expression, the induction condition was optimized at 16°C with 0.1 mM of IPTG. These induction conditions were different from those used for purifying the catalytic region of rabbit PTP-oc [13],
and from those used for purifying most PTPs [32,37,38]. E. Coli cells were harvested and broken up after 24 h of induction, and total cell lysates were analyzed by SDS-PAGE (Fig. 2). A band at the expected molecular mass was detected and reached a high level of expression 24 h after induction. As Fig. 2 shows, this band was absent in the non-induced BL21 (DE3) lysate. Because the recombinant protein was highly overexpressed under these conditions, we established an induction time of 24 h for the large-scale production of ∆PTP-oc.
Fig. 2. Recombinant ∆PTP-oc protein expression by SDS-PAGE. MW, protein marker. Lane 1, E. coli cell lysate insoluble fraction without induction. Lane 2, E. coli cell lysate insoluble fraction induced with 0.1 mM IPTG at 16°C. Lane 3, E. coli cell lysate soluble fraction without induction. Lane 4, E. coli cell lysate soluble fraction induced with 0.1 mM IPTG at 16°C. Lane 5, the purified ∆PTP-oc protein.
Purification of His6-tagged ∆PTP-oc His6-∆PTP-oc in crude extract was successfully bound to and competitively eluted from a metal-chelating affinity column after inclusion of a high concentration of imidazole (0.2 M). In the equilibration buffer, a low concentration of imidazole (10 mM) reduced the nonspecific binding of non-tagged proteins, and 500 mM NaCl prevented undesirable ionic interactions. The purification yielded a significant amount of protein, with considerable enrichment of ∆PTP-oc. Peak fractions of proteins from the chromatographic separation were resolved by SDS-PAGE and showed approximately 80% purity (Fig. 2).The identity of ∆PTP-oc was also confirmed by western blot analysis with an anti-His6 tag monoclonal antibody (Fig. 3). The molecular sizes of the recombinant proteins on SDS gels matched those predicted from the primary structures [13]. The efficiencies of the affinity purifications are shown in Table 1. The total protein yield was approximately 7 mg per 400 ml of cell culture.
R Western Marker Trans™; Lane 2, Fig. 3. Western blot analysis of recombinant ∆PTP-oc. Lane 1, EasySee○
western blot analysis of purified ∆PTP-oc with an anti-His6 tag monoclonal antibody and the protein amount was 2 µg.
Table 1 Purification of His6-tagged ∆PTP-oc*. Fractions
Total protein (mg)
Specific activity (U/mg)
Total activity (U)
Cytosolic extract
181
2.32 × 103
4.20 × 105
4
5
NIA-agarose
7
1.57 × 10
1.10 × 10
Overall yield (%)
Purification (fold)
100
1
26
6.8
*PTP activity was analyzed using 10 mM p-NPP at pH 7.0. The protein concentrations were determined using the Bradford method, with BSA as a reference standard. The volume of cell culture was 400 ml.
Phosphatase activity of ∆PTP-oc Because previous studies showed that PTPs are significantly active against p-NPP in vitro, the enzyme activity of ∆PTP-oc was assessed by measuring the ability of the protein to hydrolyze p-NPP [32,37,38]. The purified ∆PTP-oc was subjected to the PTP assay and showed highly efficient phosphatase activity for dephosphorylating p-NPP in a dose-dependent manner (Fig. 4). ∆PTP-oc also showed a time-dependent increase in the release of p-nitrophenol until the end of the reaction (Fig.4). Based on these results, the mean ∆PTP-oc specific activity was calculated to be 1.57× 104 U/mg, and was similar to those reported PTPs [37-40], suggesting that ∆PTP-oc maintains its native structure and remains intact after purification (Table 1).
Fig. 4. PTP assay of His6-∆PTP-oc. Purified recombinant His6-∆PTP-oc was assayed with two enzyme concentrations (0.1 µg/µl and 0.05 µg/µl) and different incubation times (5–30 min) in triplicate by use of p-NPP as a substrate. The relative activity is expressed as A405. Samples without enzymes were used as negative controls.
Characterization of ∆PTP-oc In general, most characterized PTPs and dual-specificity protein phosphotases display an optimal pH of approximately 5.0 when using low-molecular weight compounds as substrates [32,38,39]. However, ∆PTP-oc displayed an optimum pH of 7.0 when using p-NPP as a substrate (Fig. 5A). We also discovered a major effect on the activity at different temperatures (Fig. 5B). When the temperature was approximately 34°C, ∆PTP-oc showed maximum activity. At 37°C, the activity was 91.42% of that obtained at the optimal temperature. It was observed that changes in ionic strength had a conspicuous effect on the activity on ∆PTP-oc (Fig. 5C).
Fig. 5. Effects of pH, temperature and ionic strength on the activity of ∆PTP-oc. The activity of ∆PTP-oc was analyzed with 10 mM p-NPP at different pH values and temperatures and in the absence of NaCl. (A) The buffers used were 25 mM sodium acetate (pH 3.5–5.5), 25 mM MOPS-NaOH (pH 5.5–7.5), and 25 mM Na2B 4O7· 10H2O-HCl (pH 8.0–9.0). The reaction temperature was 37°C. (B) In the absence of NaCl, 25 mM MES-NaOH (pH 7.0) was used in the p-NPP hydrolysis assay to test the effects of diffierent temperature. (C) The effect of ionic strength on enzyme activity was tested in the presence of the indicated concentrations of NaCl at pH 7.0. The data represent the relative activities.
Kinetics of ∆PTP-oc We characterized the kinetics of ∆PTP-oc using p-NPP at pH 7.0. The data were analyzed with Graphpad Prism 4.0. The value of Vmax and Km of the enzyme with p-NPP as the substrate were estimated to be 6.1 µM/min and 801.8 µM, respectively (Fig. 6). In this case, ∆PTP-oc displayed low Km and high calculated Vmax, which indicates that ∆PTP-oc is a high efficient enzyme.
Fig. 6. Enzymatic kinetics of ∆PTP-oc. PTP activity assays were performed with different concentrations of p-NPP at pH 7.0, and the reaction velocity was calculated. (Reaction velocity=A405 /[1.78 × 104 M-1 cm-1 × reaction time]).
Conclusions Protein tyrosine phosphotases comprise a diverse super-family of enzymes that have fundamental roles in many biological processes including cell division, differentiation and gene expression [1,3,26]. PTPs are widely expressed in various cells, but their functions remain largely unknown [1,26,41]. It has been reported that rabbit osteoclastic PTP is expressed in the precursors of osteoclasts and has a positive effect on their functions [13,15,16,21,22,24,25]. In the present study, we achieved the soluble recombinant expression, purification and characterization of the intracellular phosphatase domain of PTP-oc. We found that responses of ∆PTP-oc to temperature and ionic strength were similar to those of other PTPs. The optimal pH value of ∆PTP-oc is approximately 7.0, unlike other PTPs, whose optimal pH values are approximately 5.0. It is indicated that PTP-oc may play a different role in biological processes. This study provides a set of expression constructs that could serve in further structural and functional studies.
Acknowledgments This work was sponsored by the National Natural Science Foundation of China (grant No. 811709999/H1409) and the Research Fund for the National Natural Science Foundation of Jilin Province (No. 2011-15108).
Abbreviations used: PTP-oc, osteoclastic protein tyrosine phosphatase; PTP, protein tyrosine phosphatase; ∆PTP-oc, the intracellular catalytic domain of PTP-oc; PTKs, protein tyrosine kinases; PCR, polymerase chain reaction; LB, Luria-Bertani; IPTG, isopropyl-β-Dthiogalactopyranoside; SDS-PAGE, sodium dodecyl sulfatepolyacrylamide gel electrophoresis; p-NPP, para-nitrophenylphosphate.
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Successful soluble expression of catalytic domain of PTP-oc in E. coli cells. We purified the His6-∆PTP-oc with a high purity. ∆PTP-oc exhibited classical Michaelis-Menten kinetics. ∆PTP-oc displayed an optimum pH of 7.0 At 34°C, ∆PTP-oc showed maximum activity.
S1046-5928(14)00269-1 http://dx.doi.org/10.1016/j.pep.2014.11.008 YPREP 4604
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Protein Expression and Purification
Received Date: Revised Date:
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Please cite this article as: H. Jiang, S. Yuan, C. Yue, P. Lin, W. Li, S. Xing, D. Wang, M. Hu, X. Fu, Expression, purification, and characterization of human osteoclastic protein-tyrosine phosphatase catalytic domain in Escherichia coli, Protein Expression and Purification (2014), doi: http://dx.doi.org/10.1016/j.pep.2014.11.008
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Expression, purification, and characterization of human osteoclastic protein-tyrosine phosphatase catalytic domain in Escherichia coli Huan Jianga, Sui Yuanb, Cui Yuea, Peng Lina,Wannan Lib, Shu Xingb, Deli Wangc, Min Hua*, Xueqi Fub*
a
Department of Orthodontics, School of Stomatology, Jilin University, 1500 Qinghua Road, Changchun 130021, P. R. China. b
Edmond H. Fischer Signal Transduction Laboratory, School of Life Sciences, Jilin University, Changchun 130012, P. R. China. c
School of Life Sciences, Jilin University, Changchun 130012, P. R. China.
The authors report no commercial, proprietary, or financial interest in the products or companies described in this article. Supported by the National Natural Science Foundation of China (811709999/H1409).
*Corresponding author: Min Hu, Department of Orthodontics, College of Stomatology, Jilin University. No. 1500, Qinghua Rd, Changchun 130021, Jilin, P. R. China; E-mail: [email protected]. Tel.: +86-431-887-96023; Fax: +86-431-889-75348.
Xueqi Fu, Edmond H. Fischer Signal Transduction Laboratory, College of Life Sciences, Jilin University, No. 2699, Qianjin Street,Changchun 130021, Jilin, P. R. China; E-Mail: [email protected]. Tel.: +86-431-851-55321; Fax: +86-431-851-55152.
ABSTRACT Osteoclastic protein tyrosine phosphatase (PTP-oc) is a structurally unique transmembrane protein tyrosine phosphatase (PTP) that contains only a relatively small intracellular PTP catalytic domain, does not have an extracellular domain, and lacks a signal peptide proximal to the NH2 terminus. The present study reports the expression, purification, and characterization of the intracellular catalytic domain of PTP-oc (∆PTP-oc). ∆PTP-oc was expressed in Escherichia coli cells as a fusion with a six-histidine tag and was purified via nickel affinity chromatography. When using para-nitrophenylphosphate (p-NPP) as a substrate, ∆PTP-oc exhibited classical Michaelis-Menten kinetics. Its responses to temperature and ionic strength were similar to those of other PTPs. The optimal pH value of ∆PTP-oc is approximately 7.0, unlike other PTPs, whose optimal pH values are approximately 5.0.
Keywords: Osteoclastic protein tyrosine phosphatase, Escherichia coli, Nickel affinity chromatography, Enzyme kinetics
Introduction Intracellular tyrosyl-phosphorylation levels are tightly regulated by protein tyrosine kinases (PTKs) and protein tyrosine phosphotases (PTPs) [1]. In this process, the phosphorylation reaction is mediated by PTKs, whereas the dephosphorylation reaction is catalyzed by PTPs [1,2]. The importance of tyrosine phosphorylation is illustrated best in osteoclasts [6,7], when considering the major signaling pathways, such as those mediated by RANKL and M-CSF [8,9]. Molecular studies indicate that phosphorylation of proteins on tyrosine residues is critical for the production and function of osteoclasts [2,3,4,5]. However, beyond these roles, compelling evidence also supports protein tyrosine phosphorylation having essential regulatory functions in integrin signaling in osteoclasts [10-12]. Osteoclastic protein tyrosine phosphatase (PTP-oc) belongs to the receptor PTP family and is mainly expressed in precursors of osteoclasts, including B lymphocytes, cells of the monocyte-macrophage lineage, and mature osteoclasts; it is also an activator of osteoclast differentiation and activity [13-16,20,21]. PTP-oc is the product of a gene assigned to human chromosome 12p12-p13 [17,18] or mouse chromosome 6 [19], and the expression of PTP-oc is driven by an intronic, tissue-specific proximal promoter of the PTPRO gene [13,14]. Unlike conventional transmembrane PTPs, PTP-oc is constitutive and consists of only 405 amino acid residues, which is relatively small for a transmembrane protein. The structure of PTP-oc is unique in that it contains only a single intracellular PTP catalytic domain, has a very short extracellular domain and lacks a signal peptide proximal to the NH2 terminus [13,16]. PTP-oc is an interesting protein tyrosine phosphatase, as studies show that it not only displays tissue-specific expression but also has tissue-specific functions [14]. PTP-oc may function as a positive regulator of osteoclast activity, proliferation, and differentiation [16,21 ]. Upon targeted overexpression of wildtype PTP-oc, osteoclast-like cells derived from human monocytic U937 cells created resorption pits that were larger and deeper than those of osteoclast-like cells derived from cells overexpressing phosphatase-deficient PTP-oc [16]. More importantly, more significant trabecular bone loss occurred because of increased bone resorption, but not decreased bone formation, in the osteoclastic lineage overexpressing PTP-oc in vivo [15,16, 20]. PTP-oc
stimulates osteoclasts in part by activating c-Src through dephosphorylation of its Tyr-527 (pY527) residue [15,16, 22-24]. The suppression of PTP-oc expression by a PTP-oc antisense oligodeoxynucleotide in osteoclasts reduced their bone resorption activity and increased their c-Src pY527 level [15,24]. Collectively, these results suggest that PTP-oc dephosphorylates and activates c-Src in osteoclasts; therefore, the downstream effects of this signaling may be mediated via activation of NFêB and JNK2 [22]. In addition to its regulation of osteoclast activity, there is evidence that PTP-oc may regulate osteoclast differentiation by RANKL in vitro [1,25]. Therefore, we highlight the essential regulatory role that PTP-oc may play in osteoclast function. To further study the molecular mechanism of PTP-oc signal transduction, we established an expression and purification system that is capable of providing a large-scale quantity of the functional intracellular catalytic domain of PTP-oc (∆PTP-oc). We cloned the cDNA of ∆PTP-oc into the pET-28a (+) vector, purified the His6-tagged recombinant proteins using nickel affinity chromatography, and performed detailed characterization.
Materials and Methods Construction of expression vector The pET-28a (+) plasmid was used to clone the ∆PTP-oc gene fragment. The cDNA fragment encoding the ∆PTP-oc (nucleotides 870–1656) was amplified by polymerase chain reaction (95°C for 3 min; 94°C for 1 min; 54°C for 1 min; 72°C for 1 min; 72°C for 10 min; 30 cycles) from a human cDNA library (NCBI gene ID: NM_030669.2) and was ligated into the pET-28a (+) vector by T4-DNA ligase (New England Biolabs). The primers were designed to create NcoI and XhoI restriction sites to facilitate the insertion of the ∆PTP-oc fragment into the pET-28a (+) vector. The forward primer was 5′- AGGCGCCATGGGCAAATTTTCTCTTCAGTTTGAGG -3′, and the reverse primer was 5′- GCGATCCTCGAGCACACACTGATGGATAAAAATG -3′. The ligated samples contained a C-terminal nucleotide sequence coding for six histidines, which was placed downstream and in frame with the ∆PTP-oc insert. The sizes of the polymerase chain reaction (PCR) products were confirmed by electrophoresis in a 2% agarose gel. A single colony obtained on Luria-Bertani (LB)-kanamycin plates was selected and cultured. The palsmids were purified and the sequence was confirmed by DNA sequencing. The recombinant plasmids were transformed into BL21 (DE3) cells for protein expression. Expression of His6-tagged ∆PTP-oc BL21 (DE3) cells that harbored the plasmid containing the ∆PTP-oc insert were grown in 2 ml of LB medium supplemented with 50 µg/ml kanamycin. After overnight growth at 37°C with shaking, 2 ml of this pre-culture was used to inoculate 200 ml of fresh LB media containing an appropriate antibiotic in a flask, and the cells were grown at 37°C to mid-log phase. Isopropyl-β-D- thiogalactopyranoside (IPTG) was added at a final concentration of 0.1 mM to induce ∆PTP-oc expression, and the temperature was decreased to 16°C to improve the soluble expression level [30]. After 24 h of growth at 16°C, the cells were harvested at 4°C by centrifugation at 5000 g. The pellets were resuspended in 10 ml/g extraction buffer (50 mM Tris-HCl, pH 7.4, 50 mM NaCl, 10 mM imidazole, 10% glycerol, 20 mM 2-mercaptoethanol, 2% Tween-20) containing a protease inhibitor mixture (1 mM benzamidine, 0.1 mM phenylmethylsulfonyl fluoride, 1 mM pepstatin A, 2 µg/ml leupeptin and 1 µg/ml aprotinin) and broken up by sonication on ice. The lysate was cleared by centrifugation at 13,000 g at 4°C for 30
min and both supernatant and pellets were collected. The expression levels and solubility of ∆PTP-oc were assessed by 12% sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE).
Purification of ∆ PTPPTP -oc The cytosolic supernatant from the above-mentioned step was loaded onto a Ni-NTA agarose column equilibrated with buffer A (50 mM Tris-HCl, pH 7.4, 50 mM NaCl, 10 mM imidazole, 10% glycerol). The column was washed with buffer B (50 mM Tris-HCl, pH 7.4, 500 mM NaCl, 10 mM imidazole, 10% glycerol). The tightly bound proteins were eluted with a linear imidazole gradient of 0–0.2 mol/L, and the protein peak was collected in fractions. The purity of soluble His6-∆PTP-oc was observed on 12% SDS gels. All purification procedures were carried out at 4°C and PTP activity was analyzed by use of para-nitrophenylphosphate (p-NPP) as a substrate.
Protein concentration measurements and SDSSDS-PAGE The protein concentrations were determined using the Bradford method with bovine serum albumin (Sigma) as a reference standard according to the manufacturer’s instructions [31]. Soluble His6-∆PTP-oc was concentrated to 1 mg/ml and separated on 12% Tris-glycine SDS gels. Electrophoresis was performed in a Gibco-BRL Vertical Cell Electrophoresis Apparatus (Life Technologies) with standard molecular mass markers (Transgen Biotech). The gels were stained with Coomassie Blue for 1 h at room temperature. The destaining was performed in 10% acetic acid.
Western blot assays The proteins were separated by 12% SDS-PAGE and then transferred to polyvinylidene difluoride (PVDF) membranes. Western blot analysis was performed by use of an anti-His6 tag monoclonal antibody (Abcam) as primary antibody. Proteins were detected using the enhanced chemiluminescence (ECL) method.
Enzymatic Characterization of ∆ PTPPTP -oc The phosphatase activity of the His6-∆ PTP-oc fusion protein was tested in vitro by p-NPP (Sigma) hydrolysis assays at neutral pH, as previously described [13,27-29]. The reaction mixture consisted of 25 mM MOPS-NaOH (pH 7.0), 1 mM EDTA, 1 mM dithiothreitol, 0.1 mg/ml BSA, 0.05µg/µl purified proteins and p-NPP at the indicated concentrations in a final volume of 100 µl. All activity assays were tested in triplicate. The blanks were prepared in the same manner as the samples but without the enzyme. The reaction was carried out at 37°C for 15 min and terminated by the addition of 100 µl 0.2 M NaOH solution, and the assay results were read against a blank at 405 nm. One unit of activity (U) was defined as 1 nmol of phosphate released per min, and the specific activity was expressed as U/mg [32]. Kinetics of ∆PTP-oc To study the enzyme kinetics of ∆PTP-oc, we performed enzyme assays with various concentrations of p-NPP substrate (S) for 15 min at 37 °C and pH 7.0. The data were processed with Graphpad Prism 4.0.
Results and discussion
Construction of an expression plasmid encoding ∆ PTPPTP-oc with a His tag Primary structure analyses suggest that PTP-oc is a transmembrane protein [13,14,16]. The expression of the full-length enzyme resulted in the distribution of most of the protein in inclusion bodies with relatively little PTP activity in Escherichia coli [32]. This result made the purification of active PTP-oc from E. coli cells unfeasible. The strategy to clone and express recombinant ∆PTP-oc involved inserting the gene of interest into the pET-28a (+) expression vector. Specifically, the insert encoding the phosphatase domain of PTP-oc, which was successfully amplified from the full-length human PTP-oc cDNA from pMD18-T, with the NcoI and XhoI restriction sites, was cloned into the pET-28a (+) vector containing a C-terminal His6 tag to generate pET28a-∆PTP-oc (Fig. 1A). Further characterization using specific restriction endonuclease digests and DNA sequencing demonstrated that the amplified target gene was inserted into the correct open reading frame of the vector (Fig. 1B). The resultant expression construct produced a fusion protein comprising ∆PTP-oc and a LEHHHHHH tag at the C-terminus with a predicted molecular weight of 31.2 kDa, for protein purification by metal affinity chromatography. The ligation products were then transformed into BL21(DE3) competent cells.
Fig. 1. Cloning of pET28a-△PTP-oc. (A) Cloning of pET28a-△PTP-oc from a human cDNA library (NCBI gene ID: NM_030669.2). Lane 1, Molecular weight marker DL-6000; Lane 2, △PTP-oc cloning product. (B) The double digestion of pET28a-△PTP-oc by NcoI and XhoI. Lane 1, Molecular weight marker DL-15000; Lane 2, pET28a-△PTP-oc plasmid; Lane 3, pET28a-△PTP-oc cut by NcoI and XhoI.
Expression of ∆ PTPPTP -oc in E. E . coli The relationship between the temperature dependence of protein induction and inclusion body formation has been extensively studied in a variety of model systems [33,34]. Indeed, an apparent correlation has been noted. Whereby a decrease in the induction temperature gave rise to a lower propensity for recombinant protein expression in inclusion bodies and increased the solubility of recombinant PTPs [33,35,36]. Different temperatures were attempted after induction: 37°C, 32°C, 26°C, 20°C and 16°C. The temperature was finally lowered to 16°C post-induction which highly increased the soluble expression level. After determining the optimal conditions for ∆PTP-oc expression, the induction condition was optimized at 16°C with 0.1 mM of IPTG. These induction conditions were different from those used for purifying the catalytic region of rabbit PTP-oc [13],
and from those used for purifying most PTPs [32,37,38]. E. Coli cells were harvested and broken up after 24 h of induction, and total cell lysates were analyzed by SDS-PAGE (Fig. 2). A band at the expected molecular mass was detected and reached a high level of expression 24 h after induction. As Fig. 2 shows, this band was absent in the non-induced BL21 (DE3) lysate. Because the recombinant protein was highly overexpressed under these conditions, we established an induction time of 24 h for the large-scale production of ∆PTP-oc.
Fig. 2. Recombinant ∆PTP-oc protein expression by SDS-PAGE. MW, protein marker. Lane 1, E. coli cell lysate insoluble fraction without induction. Lane 2, E. coli cell lysate insoluble fraction induced with 0.1 mM IPTG at 16°C. Lane 3, E. coli cell lysate soluble fraction without induction. Lane 4, E. coli cell lysate soluble fraction induced with 0.1 mM IPTG at 16°C. Lane 5, the purified ∆PTP-oc protein.
Purification of His6-tagged ∆PTP-oc His6-∆PTP-oc in crude extract was successfully bound to and competitively eluted from a metal-chelating affinity column after inclusion of a high concentration of imidazole (0.2 M). In the equilibration buffer, a low concentration of imidazole (10 mM) reduced the nonspecific binding of non-tagged proteins, and 500 mM NaCl prevented undesirable ionic interactions. The purification yielded a significant amount of protein, with considerable enrichment of ∆PTP-oc. Peak fractions of proteins from the chromatographic separation were resolved by SDS-PAGE and showed approximately 80% purity (Fig. 2).The identity of ∆PTP-oc was also confirmed by western blot analysis with an anti-His6 tag monoclonal antibody (Fig. 3). The molecular sizes of the recombinant proteins on SDS gels matched those predicted from the primary structures [13]. The efficiencies of the affinity purifications are shown in Table 1. The total protein yield was approximately 7 mg per 400 ml of cell culture.
R Western Marker Trans™; Lane 2, Fig. 3. Western blot analysis of recombinant ∆PTP-oc. Lane 1, EasySee○
western blot analysis of purified ∆PTP-oc with an anti-His6 tag monoclonal antibody and the protein amount was 2 µg.
Table 1 Purification of His6-tagged ∆PTP-oc*. Fractions
Total protein (mg)
Specific activity (U/mg)
Total activity (U)
Cytosolic extract
181
2.32 × 103
4.20 × 105
4
5
NIA-agarose
7
1.57 × 10
1.10 × 10
Overall yield (%)
Purification (fold)
100
1
26
6.8
*PTP activity was analyzed using 10 mM p-NPP at pH 7.0. The protein concentrations were determined using the Bradford method, with BSA as a reference standard. The volume of cell culture was 400 ml.
Phosphatase activity of ∆PTP-oc Because previous studies showed that PTPs are significantly active against p-NPP in vitro, the enzyme activity of ∆PTP-oc was assessed by measuring the ability of the protein to hydrolyze p-NPP [32,37,38]. The purified ∆PTP-oc was subjected to the PTP assay and showed highly efficient phosphatase activity for dephosphorylating p-NPP in a dose-dependent manner (Fig. 4). ∆PTP-oc also showed a time-dependent increase in the release of p-nitrophenol until the end of the reaction (Fig.4). Based on these results, the mean ∆PTP-oc specific activity was calculated to be 1.57× 104 U/mg, and was similar to those reported PTPs [37-40], suggesting that ∆PTP-oc maintains its native structure and remains intact after purification (Table 1).
Fig. 4. PTP assay of His6-∆PTP-oc. Purified recombinant His6-∆PTP-oc was assayed with two enzyme concentrations (0.1 µg/µl and 0.05 µg/µl) and different incubation times (5–30 min) in triplicate by use of p-NPP as a substrate. The relative activity is expressed as A405. Samples without enzymes were used as negative controls.
Characterization of ∆PTP-oc In general, most characterized PTPs and dual-specificity protein phosphotases display an optimal pH of approximately 5.0 when using low-molecular weight compounds as substrates [32,38,39]. However, ∆PTP-oc displayed an optimum pH of 7.0 when using p-NPP as a substrate (Fig. 5A). We also discovered a major effect on the activity at different temperatures (Fig. 5B). When the temperature was approximately 34°C, ∆PTP-oc showed maximum activity. At 37°C, the activity was 91.42% of that obtained at the optimal temperature. It was observed that changes in ionic strength had a conspicuous effect on the activity on ∆PTP-oc (Fig. 5C).
Fig. 5. Effects of pH, temperature and ionic strength on the activity of ∆PTP-oc. The activity of ∆PTP-oc was analyzed with 10 mM p-NPP at different pH values and temperatures and in the absence of NaCl. (A) The buffers used were 25 mM sodium acetate (pH 3.5–5.5), 25 mM MOPS-NaOH (pH 5.5–7.5), and 25 mM Na2B 4O7· 10H2O-HCl (pH 8.0–9.0). The reaction temperature was 37°C. (B) In the absence of NaCl, 25 mM MES-NaOH (pH 7.0) was used in the p-NPP hydrolysis assay to test the effects of diffierent temperature. (C) The effect of ionic strength on enzyme activity was tested in the presence of the indicated concentrations of NaCl at pH 7.0. The data represent the relative activities.
Kinetics of ∆PTP-oc We characterized the kinetics of ∆PTP-oc using p-NPP at pH 7.0. The data were analyzed with Graphpad Prism 4.0. The value of Vmax and Km of the enzyme with p-NPP as the substrate were estimated to be 6.1 µM/min and 801.8 µM, respectively (Fig. 6). In this case, ∆PTP-oc displayed low Km and high calculated Vmax, which indicates that ∆PTP-oc is a high efficient enzyme.
Fig. 6. Enzymatic kinetics of ∆PTP-oc. PTP activity assays were performed with different concentrations of p-NPP at pH 7.0, and the reaction velocity was calculated. (Reaction velocity=A405 /[1.78 × 104 M-1 cm-1 × reaction time]).
Conclusions Protein tyrosine phosphotases comprise a diverse super-family of enzymes that have fundamental roles in many biological processes including cell division, differentiation and gene expression [1,3,26]. PTPs are widely expressed in various cells, but their functions remain largely unknown [1,26,41]. It has been reported that rabbit osteoclastic PTP is expressed in the precursors of osteoclasts and has a positive effect on their functions [13,15,16,21,22,24,25]. In the present study, we achieved the soluble recombinant expression, purification and characterization of the intracellular phosphatase domain of PTP-oc. We found that responses of ∆PTP-oc to temperature and ionic strength were similar to those of other PTPs. The optimal pH value of ∆PTP-oc is approximately 7.0, unlike other PTPs, whose optimal pH values are approximately 5.0. It is indicated that PTP-oc may play a different role in biological processes. This study provides a set of expression constructs that could serve in further structural and functional studies.
Acknowledgments This work was sponsored by the National Natural Science Foundation of China (grant No. 811709999/H1409) and the Research Fund for the National Natural Science Foundation of Jilin Province (No. 2011-15108).
Abbreviations used: PTP-oc, osteoclastic protein tyrosine phosphatase; PTP, protein tyrosine phosphatase; ∆PTP-oc, the intracellular catalytic domain of PTP-oc; PTKs, protein tyrosine kinases; PCR, polymerase chain reaction; LB, Luria-Bertani; IPTG, isopropyl-β-Dthiogalactopyranoside; SDS-PAGE, sodium dodecyl sulfatepolyacrylamide gel electrophoresis; p-NPP, para-nitrophenylphosphate.
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Successful soluble expression of catalytic domain of PTP-oc in E. coli cells. We purified the His6-∆PTP-oc with a high purity. ∆PTP-oc exhibited classical Michaelis-Menten kinetics. ∆PTP-oc displayed an optimum pH of 7.0 At 34°C, ∆PTP-oc showed maximum activity.
