High-level expression of prolyl endopeptidase in Pichia pastoris using PLA2 as a fusion partner

Journal of Molecular Catalysis B: Enzymatic 125 (2016) 81–87

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High-level expression of prolyl endopeptidase in Pichia pastoris using PLA2 as a fusion partner Ting Jiang a , Chao Kang a,b , Xiao-Wei Yu a,∗ , Yan Xu a,∗ a The Key Laboratory of Industrial Biotechnology of Ministry of Education, State Key Laboratory of Food Science and Technology, School of Biotechnology, Jiangnan University, 1800 Lihu Ave, Wuxi, 214122 Jiangsu, PR China b Research Institute of Food Science & Engineering Technology, Hezhou university, 18 Xihuan Road, Hezhou, 542899 Guangxi, PR China

a r t i c l e

i n f o

Article history: Received 13 November 2015 Received in revised form 11 January 2016 Accepted 11 January 2016 Available online 15 January 2016 Keywords: Prolyl endopeptidase Fusion protein PLA2 High expression Properties

a b s t r a c t In our previous studies, a prolyl endopeptidase (PEP) gene from Aspergillus oryzae (MOH) was cloned and expressed in Pichia pastoris; however, the recombinant protein expression level of MOH was very low. In the present study, the PEP expression level was successfully improved by constructing fusion expression proteins with four fusion partners, namely, Streptomyces violaceoruber Phospholipase A2 (PLA2 ), cellulosebinding domain (CBD), small ubiquitin-related modifier (SUMO) and maltose binding protein (MBP). The enzyme activities of the recombinant fusion proteins CLMH, SLMH, MLMH and PLMH were increased to 3.8-, 2.7-, 4.9- and 7.4-fold compared with that of the parent MOH. Moreover, the extracellular protein content of CLMH, SLMH, MLMH, PLMH were 1.42-, 1.25-, 1.67- and 1.83-fold higher compared with that of MOH. Both PLMH and MOH showed the highest activity at pH 5.5, the highest stability at pH 6.0 and maximal activity at 40 ◦ C. © 2016 Elsevier B.V. All rights reserved.

1. Introduction Prolyl endopeptidase [EC.3.4.21.26] (PEP), also known as proline-specific endoprotease and prolyl oligopeptidase, is a special serine protease with a unique ability to cleave peptides bonds on the C-terminal side of internal proline residues [1]. Because of its ability to improve the non-biological stability of beer [2] and effectively hydrolyze haze-active protein in beer haze formation [3], PEP and its hydrolysis products are widely applied in the food manufacturing industry. PEP has also been extensively investigated as a potential pharmaceutical target for various diseases such as neurodegenerative disorder, Alzheimer’s disease [4] and learning and spatial memory impairment [5] and for the regulation of glucose metabolism and pancreatic function [6]. Therefore, obtaining

Abbreviations: PEP, prolyl endopeptidase; MOH, recombinant PEP; PLA2 , phospholipase A2 ; CBD, cellulose-binding domain; SUMO, small ubiquitin-related modifier; MBP, maltose binding protein; GST, glutathione-S-transferase; BMGY, buffered minimal glycerol-complex medium; BMMY, buffered minimal methanolcomplex medium; LB, luria-bertani; LLB, low salt luria-bertani; CLMH, CBD fused with MOH; SLMH, SUMO fused with MOH; MLMH, MBP fused with MOH; PLMH, PLA2 fused with MOH; PMF, peptide mass fingerprinting; DTT, dithiothreitol; IAM3, iodacetamide; CAN, acetonitrile. ∗ Corresponding author. Fax: +86 0510 85918201. E-mail addresses: [email protected] (T. Jiang), [email protected] (X.-W. Yu), [email protected] (Y. Xu). http://dx.doi.org/10.1016/j.molcatb.2016.01.005 1381-1177/© 2016 Elsevier B.V. All rights reserved.

high-level expression of PEP from a microorganism with enhanced activity is attractive for research and industrial applications. In 1980, a Japanese scholar found that a secreted PEP was produced by Elizabethkingia meningoseptica, which was the first PEP that was purified in a microorganism. The activity of PEP was 34.5 U/g (wet cells weight) [7]. When the E. meningoseptic PEP was expressed by Escherichia coli, its activity reached 558 U/g (wet cell weight), and the specific activity was 84 U/mg, which was much higher than that of the original strain [8]. In 1993, the PEP gene from Aeromonas hydrophila was cloned, sequenced and expressed in E. coli JM83, and the specific activity of PEP reached 320 U/g [9]. Moreover, Aspergillus fumigatus CICIM F0044 PEP cDNA was expressed in Pichia pastoris, and a maximum enzyme activity of 647.3 U/L was obtained from the recombinant yeast [10]. In summary, thus far, PEP’s yield and activity are not high. Therefore, using an effective protein expression strategy is essential for enhancing PEP secretion and accelerating its practical and industrial application. Secretory expression of heterologous proteins is often subject to several bottlenecks that limit yield, which indicates the necessity of developing a novel strategy for a secretion system. Fusion with other protein molecules or subdomains, tags and signal-anchor peptides can result in improved protein secretion [11,12]. Fusion expression has a high impact, particularly in the large-scale industrial production of heterologous proteins, where secretory expression is important for simplifying the downstream protein purification process and improvement of

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T. Jiang et al. / Journal of Molecular Catalysis B: Enzymatic 125 (2016) 81–87 Table 1 Primers used in the present study. Primers

Sequences (5 –3 )

PLA2 -F PLA2 -R CBD-F CBD-R SUMO-F SUMO-R MBP-F MBP-R PEP-F PEP-R

ATCAGAATTCGCTCCACCTCAGGCTGC ACAATCCTAAAGAACCACCACCACCAGAACCACCACCACCAAGAATTTTC AGAATTCCAGCAGACTGTCTGGGGACA AGAACCACCACCACCAATGCATTGGGCATA AGAATTCGGTCACCATCATCATCATCA AGAACCACCACCACCGGGATCAAACTCACC AGAATTCGCTAGTAAGATCGAAGAAGG AGAACCACCACCACCTCCAGCACCATCGCC GGTGGTGGTGGTTCTGGTGGTGGTGGTTCTTTAGGATTGT AGCGGCCGCCATAACTGCACCCTTAG

The linker was highlighted by bold.

In our previous studies, we found a new food-grade microorganism from Aspergillus oryzae that can produce PEP [2]. The present paper describes the efficient secreted expression of active PEP with several fusion partners, CBD, SUMO, MBP and PLA2 , in P. pastoris GS115. All of the fusion partners effectively promoted the secretion expression of PEP. The PLA2 from Streptomyces violaceoruber was successfully used in the methylotrophic yeast P. pastoris GS115 as a fusion chaperon for the first time. Furthermore, we determined the enzymatic properties of PEP. 2. Materials and methods 2.1. Reagents P. pastoris GS115 was used as a host cell. Our laboratory constructed the plasmid pPIC9K-PLA2 containing the S. violaceoruber PLA2 gene [19] (GenBank: NO.AY359866.1) and the plasmid pPIC9K-MO containing the parent A. oryzae PEP gene (MOH) [20] (GenBank: XM 001825944.2). The pPICZ␣A vector was purchased from Invitrogen, USA. The genes CBD, SUMO and MBP were optimized in accordance with the codon preference of P. pastoris and synthesized by Sangon Biotech (Shanghai). Vector pMD-19T, T4 DNA ligase, restriction enzymes and rTaq polymerase were purchased from TaKaRa (Dalian, China). The Standard Mini Plasmid I (100) and DNA Gel Extraction (100) kits were purchased from Omega. Substrate (Ala-Ala-Pro-pNA) was synthesized by Sangon Biotech (Shanghai). All other chemicals were purchased from Sinopharm Chemical Reagent. 2.2. Media

Fig. 1. a Growth curve of recombinant strains. b Protein concentration of recombinant strains. c Recombinant strain activity in the culture supernatant.

cell specific secretion titers. Furthermore, localization of the target protein and enhancement of the solubility and stability of the recombinant protein are also part of the intrinsic commercial value of fusion expression. Some of these fusion expression partners, such as maltose-binding protein (MBP) [13], cellulose-binding domain (CBD) [14,15], small ubiquitin-related modifier (SUMO) [16,17] and glutathione-S-transferase (GST) [18], have proven their values in those field. However, to date, there are no reports on the application of fusion expression to heterologous expression of PEP.

BMGY medium (1% v/v glycerol, 2% w/v peptone, 1% w/v yeast extract, 1.34% w/v yeast nitrogen base without amino acids (YNB), 4 × 10−5 % w/v biotin and 100 mM potassium phosphate buffer, pH 6.0) was used for P. pastoris growth. BMMY medium (0.5% v/v methanol, 2% w/v peptone, 1% w/v yeast extract, 1.34% w/v YNB, 4 × 10−5 % w/v biotin and 100 mM potassium phosphate buffer, pH 6.0) was used for induction. LB medium (0.5% w/v yeast extract, 1% w/v glucose, 1% w/v NaCl and 100 ␮g/mL ampicillin) and LLB medium (0.5% w/v yeast extract, 1% w/v glucose, 0.5% w/v NaCl and 25 ␮g/mL Zeocin) were used for E. coli growth. A YPD-Zeocin plate (1% w/v yeast extract, 2% w/v peptone, 2% w/v glucose, 2% w/v agar and 100 ␮g/mL Zeocin) was used for screening. 2.3. Construction of the fusion expression strain E. coli strain JM109 and P. pastoris strain GS115 were used for plasmid constructions and protein expression, respectively. The PLA2 gene was amplified from the plasmid pPIC9K-PLA2 using primers PLA2 -F (EcoRI site italics) and PLA2 -R (NotI site italics, Table 1). The PEP gene was amplified from the pPIC9K-

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MO plasmid using the primers PEP-F and PEP-R (Table 1). The two isolated products were integrated using overlap PCR. The resulting PCR fragments were digested with the restriction enzymes EcoRI and NotI, and ligated into the corresponding sites of pPICZ␣A. The ligated constructs were transformed by heat-shock into JM109 cells. After selective growth on LLB plate at 37 ◦ C overnight, colonies were inoculated in liquid LLB (no agar) at 37 ◦ C, and cultivated for 8 h for positive clone screening. Approximately 5 ␮g of DNA linearized with SacI was transformed into P. pastoris strain GS115/pPIC9K using electroporation (Multi-Copy Pichia Expression Kit, Invitrogen) and spread on a selective YPD-Zeocin plate. Positive recombinant strains GS115/pPIC9K/pPICZ␣A-PLMH were verified using PCR amplification of genomic DNA with AOX1 and target gene primers. The parent strains GS115/pPIC9K/pPICZ␣A-MOH and other fusion expression strains GS115/pPIC9K/pPICZ␣A-CLMH, GS115/pPIC9K/pPICZ␣ASLMH and GS115/pPIC9K/pPICZ␣A-MLMH were constructed in the same way. All of these recombinant genes were constructed with a 6xHis tag. The recombinant proteins are CBD fused with MOH named CLMH, SUMO fused with MOH named SLMH, MBP fused with MOH named MLMH, and PLA2 fused with MOH named PLMH. 2.4. Cultivation of P. pastoris and PEP expression The positive recombinant strains were grown on a YPD-Zeocin plate for 2–3 days. Individual colonies were inoculated into 25 mL of BMGY medium and cultivated overnight at 30 ◦ C and 200 rpm in 250-mL glass flasks. When the OD600 reached 2–6, the cells were centrifuged and resuspended in 50 mL of BMMY medium and then shaken at 28 ◦ C and 250 rpm in 500-mL glass flasks for 120 h. The culture was supplemented with 1% v/v methanol to induce the expression of fusion protein every 24 h. At the same time, samples were taken every 24 h. After centrifugation, the cell pellets and supernatant were collected for a functional assay. 2.5. Analysis of PEP activity PEP activity was spectrophotometrically assayed by the monitoring cleavage of a synthetic peptide, Ala-Ala-Pro-pNA. Briefly, 7.545 mg of substrate (Ala-Ala-Pro-pNA), was dissolved in 4 mL of 1, 4-dioxane (40%, v/v in ddH2 O) to prepare a 5 mM solution. The reaction mixture consisted of 80 ␮l of 0.1 M citrate/disodium phosphate buffer (pH 5.0), 10 ␮L of enzyme solution and 10 ␮L of substrate solution. The enzyme activity was determined by incubating at 40 ◦ C for 5 min. The reaction products were monitored spectrophotometrically at 410 nm. One International Unit of activity was defined as the quantity of enzyme that releases 1 ␮mol of p-nitroanilide per minute under the specified conditions. All experiments were conducted in triplicate. 2.6. Peptide mass fingerprinting analysis of PLMH Peptide mass fingerprinting (PMF) was analyzed using matrixassisted laser desorption/ionization time of flight (MALDI TOF) tandem mass spectrometry (MALDITOF-MS/MS). Protein samples were separated by 12% SDS-PAGE and stained by Coomassie Blue R-250. A small piece of gel on the protein band was cut and discolored with 200–400 ␮L of 100 mM NH4 HCO3 /30% acetonitrile solutions for 1 h. The supernatant was removed, and the gel was incubated in 10 ␮L of 100 mM dithiothreitol (DTT) and 90 ␮L of 100 mM NH4 HCO3 at 56 ◦ C for 30 min. The supernatant was removed and supplemented with 70 ␮L of 100 mM NH4 HCO3 and 30 ␮L of 200 mM iodacetamide (IAM3). After treatment for 20 min in a dark place, the gel was washed with 100 ␮L of 100 mM NH4 HCO3 for 15 min and 100 ␮L of 100% acetonitrile (ACN) for 5 min. The sample was dried with filter paper and was reacted with

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5 ␮L of 2.5–10 ng/␮L trypsin for 1 h at 4 ◦ C. At the end of the reaction, the sample was incubated in 20–30 ␮L of 25 mM NH4 HCO3 at 37 ◦ C for 20 h. After trypsin digestion, sample proteins were cleaved into smaller peptide fragments, and then, the peptide fragments were introduced into a mass spectrometer (Bruker Daltonics, Germany). The positive-ion mode was employed, and the mass spectrometer with the application of a spray voltage was set at 3.2 kV. The MASCOT search tool (URL, http://www.matrixscience.com) was used for identification of tryptic maps. 2.7. Purification of the fusion protein After 84 h of high-density fed-batch fermentation, the entire medium was harvested by centrifugation at 6000 × g for 30 min to remove cells. PEP was then concentrated by ultrafiltration through a 30-kDa membrane (Millipore, USA). The concentrate was brought to 60% saturation with (NH4 )2 SO4 , left for 4 h. Then precipitate was dissolved in a small volume of 20 mM phosphate buffer (pH 5.0), and dialyzed overnight. The steps were carried out 4 ◦ C unless otherwise described. The histidine-tagged PEP from the culture supernatant were purified using Ni-NTA chromatography with an ÄKTA purifier (GECo.). The purity of the protein was monitored using SDS-PAGE. 2.8. SDS-PAGE and western blot analysis All samples were normalized using the same cell concentration (OD600 ). Denaturing SDS-PAGE was performed as previously described by Laemmli [21]. Protein samples were subjected to 12% SDS-PAGE using a Mini-Protein II Cell (Bio-Rad). Proteins were stained with Coomassie bright blue and quantified using a Molecular Imaging System, with the low protein ladder (TaKaRa, China) as a standard. For western blot analysis, the proteins were separated using electrophoresis and then transferred onto a Protran nitrocellulose membrane using a Mini Trans-Blot Cell (Bio-Rad). A purified anti-His antibody raised against the His-tag of the purified PEP protein was used as the primary antibody and was diluted 1:1000 prior to application. Horseradish peroxidase conjugated goat anti-mouse IgG was diluted 1:500 as the secondary antibody. An immunoblot assay system (Bio-Rad Laboratories) was used to quantify the relative amount of protein. A One Step Yeast Active Protein Extraction Kit (Sangon, Shanghai) was used to obtain the intracellular PEP protein. 2.9. Effect of optimum temperature and pH The optimum temperature was determined by measuring purified enzyme activity in a range from 25 to 80 ◦ C at pH 5.5. Thermal stability was determined by incubating the enzyme solution at temperatures between 25 and 80 ◦ C for 120 min and analyzing the residual activity at 40 ◦ C, pH 5.5. The effect of pH on the fusion protein was determined by examining the activity of the enzyme at an optimal temperature of 40 ◦ C in various buffers, including 0.05 M citrate buffer at pH 4.5–5.5 and 0.05 M potassium phosphate buffer at pH 6.0–8.0. For pH stability, the enzyme was incubated in buffers at a pH range from 2.2 to 12.0 for 120 min at room temperature, and the residual activity was examined at pH 5.5 and 40 ◦ C. All data were represented by the relative enzyme activity and experiments were conducted in triplicate. 2.10. Enzymes kinetics The Michaelis–Menten constant (Km ) and kcat of the purified enzyme were determined using Ala-Ala-Pro-pNA as a substrate at concentrations of 1–10 mM at pH 5.5 and 40 ◦ C using the standard method. The kinetic data were calculated from a Lineweaver–Burk

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Fig. 3. SDS-PAGE analysis of GS115/pPIC9K/pPICZ␣A-PLMH. Lane M, molecular weight marker; Lane 1, culture supernatant of GS115/pPIC9K/pPICZ␣A as negative control; Lane 2–7, culture supernatant of GS115/pPIC9K/pPICZ␣A-PLMH in 24, 48, 72, 84, 96 and 120 h, respectively.

3.2. Peptide mass fingerprinting analysis of PLMH

Fig. 2. a Western blot analysis of culture supernatant. Lane M, molecular weight marker; lane 1, MOH; lane 2, CLMH; lane 3, SLMH; lane 4, MLMH; and lane 5, PLMH; Lane 6, GS115/pPIC9K/pPICZ␣A as negative control. b Western blot analysis of the intracellular proteins. Lane M, molecular weight marker; Lane 1, MOH; lane 2, SLMH; lane 3, CLMH; lane 4, MLMH; lane 5, PLMH; lane 6, GS115/pPIC9K/pPICZ␣A as a negative control.

plot with the Michaelis–Menten equation using SkanIt RE for MS 2.4.2. 3. Results 3.1. Expression of fusion PEPs in P. pastoris To compare the influence of different fusion tags on the secretory expression of PEP, we compared the extracellular secretion level and enzymatic activity of the parent with these fusion PEPs in P. pastoris. All recombinant strains contained five copies of the integrated PEP gene. Fig. 1 shows PEP enzyme activity, cell concentration (OD600 ) and protein concentration during induction by methanol. The cell growth rates of all recombinant strains were comparable during the cultivation period. As shown in Fig. 1a, all PEPs of the cell concentration (OD600 ) were almost the same. The maximum activity of MOH, CLMH, SLMH, MLMH and PLMH were 38 ± 7.2, 146 ± 5.5, 105 ± 6.5, 187 ± 6.9 and 280 ± 7.1 U/L, respectively. In particular, the secreted PEP activity of PLMH was 7.4-fold higher than that of MOH. Moreover, western blot analysis of the culture supernatant detected an increase of the fusion secretion level compared with that of MOH (Fig. 2a), and no target band of approximately 80 kDa was detected in the culture of the blank strain GS115/pPIC9K/pPICZaA. In addition, intracellular soluble proteins showed no target strip, with the exception of the protein bands of P. pastoris itself at approximately 25 kDa (Fig. 2b), which suggested that most of the fusion proteins were secreted into the culture medium. This result indicated that the increased levels of secreted PEP activity were largely due to the increase of secreted protein production (Fig. 2). Accordingly, PLMH was chosen for subsequent studies. We measured the secretion level of PLMH from 24–120 h of cultivation. The result showed an observable increase in the secretion of PEP over time (Fig. 3).

To verify the secreted PLMH by P. pastoris, the diffused bands were excised from the gel and digested using trypsin. Peptides were analyzed using MALDI-TOF-MS/MS. As shown in Fig. 3, on the SDSPAGE gel loaded with the supernatant, four separated bands, a, b, c and d, observed at 80, 21, 18, and 14 kDa were analyzed. The MASCOT database analysis showed that the band (a) contained three unique MS/MS fragments of EAAGPDGFAPVR, YYGNSTPFPISR and QALEDIPYFAR, which belong to the PEP from A. oryzae (Fig. 4a). These bands (b, c and d) were also excised from the gel, and the results showed that they all contained a unique MS/MS fragment of RIDSAFYEDMKR, which belongs to the PLA2 fusion partner from S. violaceoruber (Fig. 4b; only the result of band c is shown). These results demonstrated that the higher band at 80 kDa (shown in band a) was the cleaved product of PEP by P. pastoris. This phenomenon could be explained by the inactivity of the full-length proenzyme (proPEP) until specific proteolytic processing removes a short Nterminal extension prosequence, with processing that appears to be performed by PEP itself [22–24]. In our experiment, we speculate that the PLA2 fusion partner, shown as bands b, c and d, was cut away at the same time during the specific proteolytic processing. The migration of these three bands of b, c and d at 21, 18 and 14 kDa, respectively, were due to N-glycosylation, which was also observed when only PLA2 was expressed in P. pastoris [19]. 3.3. Properties of PEPs The recombinant PEP MOH and fusion protein PLMH were purified as described in the “Materials and methods” (Table 2). Using Ala-Ala-Pro-pNA as a substrate, the optimal pH, pH stability, optimal temperature and temperature stability were determined for each sample as described in detail in the Methods section. The optimum temperature and stability of the parent MOH and the fusion protein PLMH were nearly the same. They both showed maximal activity at 40 ◦ C (Fig. 5a) and showed high stability (over 80% activity) between 25 and 35 ◦ C after incubation for 120 min, but they were not very stable at higher temperatures. When the temperature reached 40 ◦ C, the relative activity decreased to 30% of the maximum activity, and the activity was almost lost after incubation for 120 min at 45 ◦ C. Additionally, MOH and PLMH had the same pH profile (Fig. 5b). Within the pH range of 5.5–6.5, MOH and PLMH retained more than 80% of the initial activity after incubation at 40 ◦ C for 120 min. MOH and PLMH showed the highest activity at pH 5.5, and over 60% of the maximum activity was retained between pH 5.0 and 7.0. Kinetic data for the

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Fig. 4. a MALDI-TOF-MS/MS spectra of band a. b MALDI-TOF-MS/MS spectra of band c in Fig. 3.

purified PLMH were evaluated at 40 ◦ C and pH 5.0. The Km , kcat , and kcat /Km of PLMH were 0.23 ± 0.01 mM, 112.51 ± 0.02 S−1 and 489.17 s−1 mM−1 , respectively, which was very close to the values of MOH that were determined previously [25].

4. Discussion Here, we demonstrated that expression of A. oryzae PEP in P. pastoris was markedly enhanced upon fusion to CBD, SUMO, MBP

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Table 2 Purification of the PLMH and MOH. Purification PLMH

MOH

steps

Crude extract Ultrafiltration (30 kDa) (NH4 )2 SO4 (60%), dialyzed Ni-NTA chromatography Crude extract Ultrafiltration (30 kDa) (NH4 )2 SO4 (60%), dialyzed Ni-NTA chromatography

Totalactivity(U)

Specific activity(U/mg)

Purification(-fold)

Yield(%)

560 468 98 50 700 540 112 32.4

2.9 5.8 10.2 43.2 3.2 5.8 10.2 49.3

1 2 3.5 14.9 1 1.8 3.2 15.4

100 84 17.5 8.9 100 77 16 4.6

Fig. 5. a The effect of temperature on MOH and PLMH activity and stability. The activity for optimum temperature was determined under conditions at 25–80 ◦ C and pH 5.5. The effect of temperature on enzyme stability was determined by incubating the enzyme for 120 min at temperatures of 25–80 ◦ C at pH 5.5. b The effect of pH on MOH and PLMH activity and stability. The activity was determined under conditions at pH 2.2–12.0 and 40 ◦ C. The pH stability was determined at 40 ◦ C and pH 5.5 by separately incubating the enzyme in citrate/disodium phosphate buffer (pH 2.2–8.0), Tris–HCl (pH 9.0–10.5) and glycine/NaOH (pH 11.0–12.0) for 120 min at room temperature. All experiments were conducted in triplicate.

and PLA2 as fusion partners. When fused without CBD, SUMO, MBP and PLA2 , PEP was so poorly expressed in P. pastoris that it could hardly be detected using a specific substrate, and only a faint band (nearly 80 kDa) was observed on a western blot probed with an anti-His antibody (Fig. 2a). In contrast, when fused with these tags, the expression of PEP was dramatically increased. In

particular, when fused with PLA2 , the effect was more significant. Previous work has shown that SUMO, CBD and MBP were capable of enhancing the expression level and stability of heterologous proteins. For example, SUMO dramatically enhanced the expression of a series proteins in different expression systems [16,26–28].

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The SUMO tag can function as a molecular chaperone to prevent the aggregation of intermediates and keep the folding intermediates in solution long enough to recover to its correct conformation [29]. For CLMH, the CBD has two disulfide bonds. These disulfide bonds may enhance the folding stability of the fusion protein CLMH [30]. In recent years, MBP has become a useful fusion strategy for heterologous protein expression [31,32]. The most popular model is that MBP possesses molecular chaperone characteristics. MBP contains hydrophobic residues in its ligand bonding cleft, which appear to play a central role in guiding the target proteins to their native structures. These properties are believed to allow MBP to reversibly bind to folding intermediates of its fusion partners and temporarily keep them in a conformation that prevents their self-association and aggregation [33,34]. In our previous study, the phospholipase A2 (PLA2 ) from S. violaceoruber was successfully expressed in the methylotrophic yeast P. pastoris GS115. When the PLA2 protein was expressed in P. pastoris, protein secretion could reach 12 g/L in high-cell-density fed-batch cultures (data not shown). Moreover, as a fusion partner, in some aspect, PLA2 has advantages over other fusion tags, such as a small molecular weight and a high ability to promote protein secretion expression. Thus, PEP expression was probably improved by the high secretion ability of PLA2 and, as a result, facilitated fusion protein secretion. In summary, the data provided here demonstrate that PEP fused with PLA2 had the highest expression level in P. pastoris. Our results indicate that this novel combination of a fusion protein domain fused to a poorly expressed protein of interest via a linker for in vivo processing can be applied as a general strategy to improve the yields of functionally active proteins. Acknowledgements Financial support from the National High Technology Research and Development Program of China (863 Program) (2012AA022207), 333 Project in Jiangsu Province (BRA2015316), Six Talent Peaks Project in Jiangsu Province (NY-010), the High-end Foreign Experts Recruitment Program (GDW20123200113), NSFC (20802027) and the 111 Project (111-2-06) are greatly appreciated. References [1] R. Mentlein, FEBS Lett. 234 (1988) 251–256. [2] C. Kang, X.W. Yu, Y. Xu, J. Ind. Microbiol. Biotechnol. 42 (2015) 263–272.

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[3] M. Lopez, L. Edens, J. Agric. Food Chem. 53 (2005) 7944–7949. [4] M. Morawski, K. Nuytens, T. Juhasz, U. Zeitschel, G. Seeger, E. Waelkens, L. Regal, I. Schulz, T. Arendt, Z. Szeltner, J. Creemers, S. Rossner, Neuroscience 242 (2013) 128–139. [5] G. Dagostino, J.D. Kim, Z.W. Liu, J.K. Jeong, S. Suyama, A. Calignano, X.B. Gao, M. Schwartz, S. Diano, Cereb. Cortex. 23 (2013) 2007–2014. [6] J.D. Kim, C. Toda, G. D’Agostino, C.J. Zeiss, R.J. DiLeone, J.D. Elsworth, R.G. Kibbey, O.W. Chan, B.K. Harvey, C.T. Richie, M. Savolainen, T. Myohanen, J.K. Jeong, S. Diano, Proc. Natl. Acad. Sci. U. S. A. 111 (2014) 14003. [7] T. Yoshimoto, R. Walter, D. Tsuru, J. Biol. Chem. 255 (1980) 4786–4792. [8] T. Diefenthal, H. Dargatz, V. Witte, G. Reipen, I. Svendsen, Appl. Microbiol. Biotechnol. 40 (1993) 90–97. [9] A. Kanatani, T. Yoshimoto, A. Kitazono, T. Kokubo, D. Tsuru, J. Biochem. 113 (1993) 790–796. [10] L. Ning-Huan, T. Zheng, S. Wei, Microbiol. China 40 (2008) 9–12. [11] M. Wu, W.H. Liu, G.H. Yang, D.K. Yu, D.H. Lin, H.Y. Sun, S.Q. Chen, Appl. Biochem. Biotechnol. 172 (2014) 2400–2411. [12] Y.S. Kang, J.A. Song, K.Y. Han, J. Lee, J. Biotechnol. 194 (2015) 39–47. [13] B. Daelken, R.A. Jabulowsky, P. Oberoi, I. Benhar, W.S. Wels, PLoS One 5 (2010) e14401. [14] J.C. Rotticci-Mulder, M. Gustavsson, M. Holmquist, K. Hult, M. Martinelle, Protein Expression Purif. 21 (2001) 386–392. [15] J.O. Ahn, E.S. Choi, H.W. Lee, S.H. Hwang, C.S. Kim, H.W. Jang, S.J. Haam, J.K. Jung, Appl. Microbiol. Biotechnol. 64 (2004) 833–839. [16] X.J. Wang, X.M. Wang, D. Teng, Y. Zhang, R.Y. Mao, J.H. Wang, Lett. Appl. Microbiol. 59 (2014) 71–78. [17] X. Dai, Z. Sun, R. Liang, Y. Li, H. Luo, Y. Huang, M. Chen, Z. Su, F. Xiao, Appl. Microbiol. Biotechnol. 99 (2015) 5997–6007. [18] K. Shimada, M. Nagano, M. Kawai, H. Koga, Proteomics 5 (2005) 3859–3863. [19] A. Liu, X.-W. Yu, C. Sha, Y. Xu, Appl. Biochem. Biotechnol. 175 (2015) 3195–3206. [20] K. Chao, X.W. Yu, X. Yan, J. Ind. Microbiol. Biotechnol. 42 (2014) 263–272. [21] U.K. Laemmli, Nature 227 (1970) 680–685. [22] M. Gamble, G. Kuenze, E.J. Dodson, K.S. Wilson, D. Jones, Proc. Natl. Acad. Sci. U. S. A. 108 (2011) 3536–3541. [23] F. Ruiz-Perez, J.P. Nataro, Cell Mol. Life Sci. 71 (2014) 745–770. [24] L.M. Babe, B. Schmidt, M. B.B.A-Protein. Struct, (1998) 211–219. [25] C. Kang, X.-W. Yu, Y. Xu, J. Ind. Microbiol. Biot. 41 (2014) 49–55. [26] X. Zuo, S. Li, J. Hall, M.R. Mattern, H. Tran, J. Shoo, R. Tan, S.R. Weiss, T.R. Butt, J. Struct. Funct. Genomics 6 (2005) 103–111. [27] L. Liu, J. Spurrier, T.R. Butt, J.E. Strickler, Protein Expression Purif. 62 (2008) 21–28. [28] R.J. Peroutka, N. Elshourbagy, T. Piech, T.R. Butt, Protein Sci. 17 (2008) 1586–1595. [29] E.S. Johnson, Annu. Rev. Biochem. 73 (2004) 355–382. [30] M. Linder, T.T. Teeri, J. Biotechnol. 57 (1997) 15–28. [31] Z. Li, W. Leung, A. Yon, J. Nguyen, V.C. Perez, J. Vu, W. Giang, L.T. Luong, T. Phan, K.A. Salazar, Protein Expression Purif. 72 (2010) 113–124. [32] R. Eliseev, A. Alexandrov, T. Gunter, Protein Expression Purif. 35 (2004) 206–209. [33] S. Nallamsetty, D.S. Waugh, Biochem. Biophys. Res. Commun. 364 (2007) 639–644. [34] J.D. Fox, K.M. Routzahn, M.H. Bucher, D.S. Waugh, FEBS Lett. 537 (2003) 53–57.