Comparative Characterization of Two Forms of Recombinant Human SPC1 Secreted from Schneider 2 Cells

Protein Expression and Purification 19, 113–124 (2000) doi:10.1006/prep.2000.1215, available online at http://www.idealibrary.com on

Comparative Characterization of Two Forms of Recombinant Human SPC1 Secreted from Schneider 2 Cells Jean-Bernard Denault, Claude Lazure,* Robert Day, and Richard Leduc 1 *Laboratory of Neuropeptide Structure and Metabolism, Institut de Recherches Cliniques de Montre´al, Montre´al, Que´bec H2W 1R7, Canada; and Department of Pharmacology, Faculty of Medicine, Universite´ de Sherbrooke, Sherbrooke, Que´bec J1H 5N4, Canada

Received December 2, 1999, and in revised form February 1, 2000

SPC1 (furin/PACE), an enzyme belonging to the S8 group of serine endoproteases, is a type I integral membrane protein that catalyzes the processing of a multitude of precursor proteins. We report here the use of transfected Drosophila melanogaster Schneider 2 cells to produce milligram amounts of two forms of recombinant human SPC1. In order to investigate the role of the cysteine-rich region (CRR) of SPC1, we compared the biochemical and enzymatic properties of hSPC1/714 that has the C-terminal tail and transmembrane region of the native enzyme removed with that of hSPC1/585 which had, in addition, the CRR deleted. Two stable cell lines were established. The S2-hSPC1/714 line secreted a major form of apparent molecular weight of 83 kDa and a minor form of 80 kDa whereas the S2-hSPC1/585 line secreted a single 59kDa protein. PNGase F treatment of the different forms demonstrated that the enzymes were glycosylated. Automated NH 2-terminal sequencing revealed that all purified forms resulted from processing at the expected zymogen activation site. Removal of the CRR resulted in a broadening of the enzyme’s pH range, a shift of K 0.5 for Ca 2ⴙ, and a shorter enzymatic half-life when compared to the longer form, which suggest that the CRR of hSPC1 may help in stabilizing the enzyme’s proteolytic activity. The use of this high-level expression system will meet the demand for material necessary to perform biochemical and structural studies that are needed to further our understanding of this and other SPCs at the molecular level. © 2000 Academic Press

Key Words: SPC1; furin; convertase; S2 cells; cysteine-rich region.

1 To whom reprint requests should be addressed. Fax: (819) 5645400. E-mail: [email protected].

1046-5928/00 $35.00 Copyright © 2000 by Academic Press All rights of reproduction in any form reserved.

In the secretory pathway, many biologically inactive precursor proteins are proteolytically processed at specific pairs of basic amino acids to produce biologically active peptides and proteins. Within the past decade, mammalian subtilisin-like proprotein convertases or SPCs 2 [SPC1 (furin/PACE), SPC2 (PC2), SPC3 (PC1/ PC3), SPC4 (PACE4), SPC5 (PC4), SPC6 (PC5/PC6), and SPC7 (PC7/LPC/PC8)] have been identified as the enzymes performing these cleavages (1,2). SPCs are mosaic proteins in that they are constituted of different domains, each having specific functions. Their primary structure reveals a signal peptide enabling the protein to enter the secretory pathway; a proregion necessary for proper folding and regulation of the enzymatic activity (3– 8); a catalytic domain with the Asp, His, and Ser triad; a P-domain necessary for calcium and pH regulation (9). Following the P-domain, the SPCs possess a highly variable C-terminus, some with transmembrane domains that anchor the enzyme within the membrane of various cellular compartments. SPC1 is certainly the most characterized member of the group considering that its biosynthesis, enzymatic properties, tissue distribution, intracellular localization, and trafficking have been thoroughly studied (10 –12 for review). The enzyme’s domain configuration is similar to all other SPCs but differs in that (i) a cysteine-rich region (CRR), made up of two repeated 2 Abbreviations used: t-Boc, N-tert-butyloxycarbonyl; CRR, cysteine-rich region; FBS, fetal calf serum; Hepes, N-(hydroxethyl)piperazine-N⬘-(2-ethanesulfonic acid); MCA, 4-methylcoumaryl-1-amide; KLH, keyhole limpet hemocyanin; LPC, lymphomas proprotein convertase; PACE, paired basic amino acid-converting enzyme; PC, proprotein convertase; pGlu, pyroglutamic acid; PNGase F, peptide-N-glycosidase F; S2, Schneider 2; SPC, subtilisinlike proprotein convertase; TBS, Tris-buffered saline; Tris, tris(hydroxymethyl)aminomethane.

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motifs of cysteine residues, follows the P-domain; (ii) a transmembrane domain anchors the enzyme within the cell’s trans-Golgi network; and (iii) the cytoplasmic tail directs SPC1⬘s intracellular distribution. The CRR has no known function but has sequence similarities to the CRR found in the TNF and EGF receptor family and to a Marfan syndrome allele (11). SPC1 cleaves many different types of precursor proteins into bioactive peptides. Although SPC1 acts predominantly to process precursors found intracellularly, its ability to translocate to the plasma membrane enables the enzyme to cleave exogenous proteins. Among them, bacterial toxins (i.e., anthrax toxin PA, Pseudomonas aeruginosa exotoxin A, diphtheria toxin) require correct processing at SPC1 recognition sites to exert their pathological effects. Also, numerous viral coat glycoproteins (i.e., avian influenza HA, HIV-1 gp160, measles virus F o) need such a cleavage to gain their mature state (11 and 13 for review). The implication of SPC1 in the diseases caused by activation of these precursors suggests that this convertase may be a good target for antipathogen drugs. Moreover, the possible involvement of SPC enzymes in the extracellular cleavage activation of many matrix metalloproteinases and cell surface adhesion molecules (11 for review) indicates that SPCs could play an indirect role in angiogenesis and cell growth. Thus, production of large quantities of SPC1 would be advantageous toward the development of specific applications such as the determination of SPC1⬘s crystal structure, as highthroughput screening of inhibitors, and in various biotechnological processes for the efficient and complete cleavage of target protein precursors (14). In order to generate enzymatically active SPC1, numerous expression systems have been used in the past; these include recombinant vaccinia virus (15), recombinant baculovirus-infected insect cells (16), and secretion in transgenic murine milk (17). These systems are not always easy to manipulate and lead to varying levels of expressed protein and, finally, purification of proteins of interest in their native state is sometimes compromised because of cell lysis. A system that would reconcile the high level expression of baculovirus-infected insect cells, without the need to generate recombinant viruses, and a system with continuous expression, without the disadvantages of long-term selection, would be a valuable tool for the study of convertases. Thus, the aims of this study are: (i) to assess the validity of using transfected Schneider 2 (S2) cells for the production of large quantities of recombinant human SPC1; (ii) to develop a simple purification protocol that leads to homogenous and enzymatically pure hSPC1; (iii) to determine its relatedness to the convertase expressed in other systems; and (iv) to get new biochemical insights on the possible role of the cysteine-rich region of hSPC1.

MATERIALS AND METHODS

Materials. All restriction and modification enzymes were from Amersham-Pharmacia Biotech or Promega Corp. Peptide-N-glycosidase F and neuraminidase were from Roche. The fluorogenic substrates t-Boc-ArgVal-Arg-Arg-MCA (Boc-RVRR-MCA) and pGlu-ArgThr-Lys-Arg-MCA (pERTKR-MCA) were from Bachem Bioscience. IPL-41 and insect lipid supplements were from Sigma-Aldrich. Yeastolate, L-glutamine, penicillin, streptomycin, and hygromycin B were from GibcoBRL. Hybond-C, secondary antibodies, and chemiluminescence detection kit were from AmershamPharmacia Biotech. Transfer vectors. To construct hSPC1 truncated mutants, the cDNA encoding full-length hSPC1 cloned in pGEM-7zf(⫹) (a generous gift from Dr. Gary Thomas, Vollum Institute, Portland, OR) was used as a PCR template. The hSPC1/714 mutation was generated by PCR with oligonucleotides 5⬘-gatgaggatccctctg-3⬘ and 5⬘-gaattccaccacctaaggcaggt-3⬘ that change Glu 715 (throughout the text, the amino acid position numbering refers to the preprofurin) to a stop codon. The PCR product was subcloned in pCR2.1 (Invitrogen Corp.) and nucleotide sequencing of the complete insert was performed. A BamHI insert was excised and subcloned into BamHI site of pGEM-7zf(⫹)/ hSPC1 replacing the 3⬘-end of hSPC1 by the truncated construction. Finally, an EcoRI insert was ligated into the EcoRI site of pAc5.1/V5-HisA used for transfection of S2 cells (Invitrogen Corp.). The hSPC1/585 was generated using the same strategy. The oligonucleotides used to change Gly 586 to a stop codon by PCR were 5⬘-gatgaggatccctctg-3⬘ and 5⬘-acttaactgctttctggaggtac3⬘. A NotI insert was ligated into the NotI site of pAc5.1/V5-HisA. The V5 epitope and polyhistidine purification tag from the pAc5.1/V5 HisA vector were not used because the stop codon from the original cDNA and constructions were kept intact. Cell culture and transfection. S2 cells were grown in complete DES expression medium (Invitrogen Corp.) supplemented with 10% FBS, 2 mM L-glutamine, 50 IU/mL penicillin, and 50 ␮g/mL streptomycin at a cell density between 0.5 and 20 ⫻ 10 6 cells/mL at 22°C in ambient air. S2 cells were cotransfected with the different pAc5.1 vectors and pCoHYGRO (Invitrogen Corp.) at a weight ratio of 19:1, respectively, by the calcium phosphate technique as described by the manufacturer and selected with 300 ␮g/mL hygromycin B for a period of 21 days. After removal of cell debris, cell cultures were scaled up and expression of hSPC1 was tested by enzymatic assays and Western blot analysis. Cells were adapted to serum-free IPL-41 medium containing insect cell lipid supplement, 4 g/L ultrafiltrated yeastolate, 2 mM L-glutamine, and antibiotics without hygromycin B by two successive passages in IPL-41

CHARACTERIZATION OF RECOMBINANT hSPC1 IN S2 CELLS

medium. Before initiating large-scale culture, cells were washed with IPL-41. The secreted proteins were harvested after 6 – 8 days of culture or when cell concentrations reached 30 ⫻ 10 6 cells/mL. Enzyme purification. Typically 400 –1000 mL of serum-free medium was collected after 6 – 8 days of culture in triple flask containers (NUNC). The medium was clarified by centrifugation at 5000g for 10 min and cooled on ice. All subsequent steps were carried out at 4°C. The proteins were precipitated with 536 g/L (80% saturation) ammonium sulfate and stirred on ice for at least 2 h. After collecting the precipitate following centrifugation at 8000g for 20 min, the proteins remaining in the supernatant were recovered by filtration through a 1.2-␮m GF/C glass filter (Whatman) using a vacuum pump. All protein samples were resuspended in at least 5% original volume in buffer A (20 mM Tris, pH 7.5, and 1 mM CaCl 2) containing 0.1 mM 1,10phenanthroline. Following removal of insoluble material by centrifugation at 15,000g, the sample was filtered through a 0.45-␮m filter and concentrated to 1/40 initial volume with a Centricon Plus-80 concentrator (30,000 MWCO, Millipore). A 10-mL fraction of concentrate was applied to a HR Sephacryl S-100 26/60 (Amersham-Pharmacia Biotech) gel filtration column at a flow rate of 0.5 mL/min. Fractions of 2.5 mL were collected. An aliquot from each fraction was tested for enzymatic activity (described below) and analyzed by SDS–PAGE and Western blot analysis. Fractions with enzymatic activity were pooled until the double of the original volume loaded was collected. Subsequently, this sample was purified by FPLC using a MonoQ 5/5 (Amersham-Pharmacia Biotech) anion-exchange column using a 0 –300 mM NaCl gradient in buffer A. Two-milliliter fractions were collected and analyzed for enzymatic activity, by SDS–PAGE and Western blotting. Protein levels were measured using Bradford reagent with ␥-globulin as a standard. After FPLC, the purified enzyme was kept in 0.1-mL aliquots containing 15% glycerol at ⫺80°C. Enzyme concentration was determined by Bradford colorimetric assay, using ␥-globulin as a standard protein and by assuming that all of the enzyme molecule was active. The value obtained was correlated by titration against AT-PDX, as described (18). Enzymatic assays. Enzymatic assays were carried out in 100 mM Hepes, pH 7.5, 1 mM CaCl 2, 1 mM ␤-mercaptoethanol, and varying concentrations of fluorogenic substrates, Boc-RVRR-MCA or pERTKRMCA in 0.5 mL. Assays were performed at 37°C for 30 min for detection of activity or for 1 h for kinetic analysis. Reactions were stopped by the addition of 0.5 mL of 10 mM EDTA. Reactions were calibrated with MCA and fluorescence was measured on a Hitachi F-2000 spectrofluorometer. Data were fitted into the standard

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pseudo-first-order equation or in an equation describing substrate inhibition (19), a modified form of the Michaelis–Menten equation in which K si is the binding constant for the second substrate molecule: v ⫽ V⬘关s兴/共K⬘M ⫹ 关s兴 ⫹ 关s兴 2 /K si兲

[1]

Production of anti-hSPC1 antibodies. The hSPC1 antibodies were obtained from rabbits injected with a peptide (Arg-Thr-Gln-Met-Asn-Asp-Asn-Arg-His-GlyThr-Arg-Cys) coupled to KLH (Pierce). The peptide, which encompasses amino acids 187–198 of human SPC1, exhibits a complete sequence identity to corresponding sequences of mouse, rat, hamster, and chicken SPC1. Rabbits were injected with antigen emulsified in an equal volume of TiterMax adjuvant (Sigma-Aldrich). Gel electrophoresis, Western blotting, and protein sequencing. All electrophoreses were performed on 1.0mm-thick, 8% SDS–PAGE. For Western blotting, the proteins were transferred on Hybond-C nitrocellulose membranes preequilibrated in transfer buffer (39 mM glycine, 48 mM Tris base, pH 8.3, 0.037% SDS, and 5% MeOH) for 15 min. Transfer was performed at 0.5 A for 3 h at 4°C. Following transfer, the membranes were incubated for 1 h at room temperature in 5% (w/v) nonfat skim milk diluted in TBS (20 mM Tris, pH 7.6, and 137 mM NaCl) with 0.1% Tween 20 (TBS-T) and then for 4 h with anti-SPC1 antiserum at 1:1000 dilution. The membranes were washed three times with TBS-T for 10 min and incubated for 1 h with horseradish peroxidase-coupled secondary antibodies (Amersham-Pharmacia Biotech) diluted 1:1500 in TBS-T. The membranes were washed again and the immunoreactive proteins were revealed by ECL detection. For glycosylation analysis, the samples were pretreated for 5 min at 95°C in the presence of 0.1% SDS. The sample was then diluted with 0.5% Nonidet P-40, incubated with 0.5 U of PNGase F in 50 mM Tris base, pH 6.8, 10 mM EDTA, or 0.5 U of neuraminidase in 50 mM sodium acetate, pH 5.5, 150 mM NaCl, and 5 mM CaCl 2 for 3 h at 37°C. For NH 2-terminal sequencing, 10 –20 ␮g of purified enzyme was concentrated using a Centricon-10 (Millipore), separated by SDS–PAGE, and transferred to Immobilon-P SQ membrane (Millipore) as described above. The membranes were washed several times in TBS to remove glycine and the proteins stained with Ponceau S. The stained protein bands were excised, cut in small pieces, and sequenced on an Applied Biosystems gas/liquid-phase Sequenator (Model 477 operating in gas-phase mode), directly coupled to an Applied Biosystems phenylthiohydantoinamino acids analyzer (Model 120A) using N-methylpiperidine as coupling buffer.

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FIG. 1. Schematic representation of hSPC1 along with the COOHterminal truncated mutants hSPC1/714 and hSPC1/585 used in this study. Signal peptide (black), proregion (gray), catalytic domain (clear), “P” domain (black dots), cysteine-rich domain (dashes), transmembrane domain (white dots), and cytoplasmic tail (lines) are shown.

RESULTS

Expression of hSPC1 in S2 cells. To investigate the role of the CRR of SPC1, we compared the biochemical and enzymatic properties of hSPC1/714 that has the C-terminal tail and transmembrane region of the native enzyme removed with that of hSPC1/585 which had, in addition, the CRR deleted. First, we established two cell lines by transfecting S2 cells with the cDNA encoding hSPC1/714 and hSPC1/585. Both forms were generated by introducing a translational stop codon after amino acid 714 and 585 respectively (Fig. 1). Western analysis using SPC1-specific antibodies showed that extracts of S2 cells transfected with plasmids containing the cDNAs for hSPC1/714 or hSPC1/ 585 contained a weak immunoreactive band corresponding to an 83-kDa form and an immunoreactive band corresponding to 59 kDa, respectively (Fig. 2A). When the media of these stable lines were analyzed for immunoreactive hSPC1, a strong signal for both the 83- and the 59-kDa forms was observed. As a control, wild-type S2 cells do not show any SPC1 immunoreactivity. Monitoring the enzymatic activity at various times, up to 5 days after passage, as illustrated in Fig. 2B, shows the accumulation in a nearly linear fashion of SPC1 activity as measured with the fluorogenic substrate Boc-RVRR-MCA. Net enzymatic activity was obtained by subtracting fluorescence measurements from ␤-galactosidase-expressing cells. The specific activity (expressed as ␮mol MCA released per min and per mg of proteins) in culture medium was slightly higher in the first two days of culture, but tended to stabilize as the total protein content and the cell number increased. Purification of recombinant hSPC1 enzymes. The main advantage of this expression system resides in its simplicity in achieving milligrams per liter of expression levels in serum-free medium. Optimal production yields were obtained when we harvested the medium of cells seeded at 2 ⫻ 10 6 cells/mL after 6 days. Our purification scheme basically involves three steps. The ammonium sulfate step has two purposes: it concentrates up to five times the original volume and removes

a large amount of components that may interfere with other purification steps without important loss of enzymatic activity, as depicted in Table 1. Recoveries in enzymatic activities upwards of 70 and 85% for hSPC1/ 714 and hSPC1/585, respectively, were achieved. As shown in Figs. 3A and 3B, the majority of the enzymatic activity, as well as the immunoreactive proteins, are present in the ammonium sulfate precipitate. Since S2 cells produce many proteins in the 200-kDa range, we then made use of gel filtration as the next purification step. This chromatographic step proved efficient with both SPC1 forms as it achieved a five- to sixfold purification factor while allowing a four- to five-

FIG. 2. Stable expression of hSPC1/714 and hSPC1/585 in S2 cells. (A) Western blot analysis of cell extracts and culture medium of wild-type S2 cells (left) and S2 cells expressing either hSPC1/714 (middle) or hSPC1/585 (right) after 36 h of accumulation in complete medium. A volume of 100 ␮L of suspension culture was centrifuged to separate the cells from medium. Each sample was then separated on an 8% SDS–PAGE, transferred to a Hybond-C membrane, and analyzed using an antiserum specific to the SPC1 catalytic domain. (B) The amount of enzymatic activity secreted from S2-hSPC1/714 and S2-hSPC1/585 was measured as a function of time as described under Materials and Methods using the fluorogenic substrate t-BocArg-Val-Arg-Arg-MCA. The results are shown following subtraction of background activity obtained from identical aliquots of medium of S2-␤-galactosidase-producing cells.

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CHARACTERIZATION OF RECOMBINANT hSPC1 IN S2 CELLS

TABLE 1 Representative Purification of the Recombinant hSPC1/714 and hSPC1/585

Purification step HSPC1/714 Culture medium 80% Ammonium-SO 4 precipitate Centriplus-80, 30,000 MWCO S-100 fractions MonoQ FPLC pooled fractions HSPC1/585 Culture medium 80% Ammonium-SO 4 precipitate Centriplus-80, 30,000 MWCO S-100 gel filtration MonoQ FPLC single peak (59 kDa) a b

Total volume (mL)

Total protein (mg)

Total activity (U) a,b

Specific activity (U/mg)

Yield (%)

Cumulative purification factor (fold)

464.0 82.0 27.5 32.9 12.0

45.5 38.0 25.4 5.18 0.59

766 538 545 445 153.7

16.8 14.5 21.5 85.9 260.5

— 70.2 71.1 58.1 20.1

— 0.9 1.3 5.1 15.5

483.0 66.0 10.3 10.6 2.85

46.0 26.4 20.1 3.42 0.43

290 249 143 124 82.6

6.3 9.4 7.1 36.3 192.1

— 85.9 57.4 42.8 28.5

— 1.5 1.1 5.8 30.5

Enzymatic assays were performed as described under Materials and Methods. One unit is the amount of enzyme that generates 1 nmol/min of MCA in a standard assay containing 50 ␮M t-boc-RVRR-MCA.

fold increase in specific activity, as indicated in Table 1. As depicted in Fig. 3C, the fractionation by gel filtration chromatography resulted in a single peak of enzymatic activity. The amount of enzymatic activity for either form is in good agreement with the amount of proteins being detected by Western blotting (data not shown). Finally, the third step used anion-exchange chromatography on a MonoQ column. Figure 3D shows the elution of proteins from the MonoQ column as a function of increasing NaCl concentration. The major peak of absorbance corresponded to the desired protein as shown by the activity found in each fraction using the Boc-RVRR-MCA substrate; this activity corresponds to SPC1 immunoreactivity, as detected by Western blotting (data not shown). The SPC1 originating from the S2-hSPC1/714 line eluted in two distinct active fractions (fractions 14 and 15). When these were individually analyzed by Western blotting, a single immunoreactive form of 83 kDa was observed in fraction 14 whereas fraction 15 had a 1:1 ratio of the 83- and 80-kDa forms (data not shown). However, in order to minimize losses, the enzymatic activity found in both fractions was combined and kept for further analysis. Indeed, analysis of the pooled sample, as seen in Fig. 4A (protein stain) and Fig. 4B (Western blot), revealed that hSPC1/714 is present as a doublet of 83 and 80 kDa with an estimated purity to be ⱖ80% by Coomassie blue and densitometry analysis. The hSPC1/585 cell line gave a single 59 kDa band that was estimated to be ⱖ90% pure. The overall recovery in terms of total enzymatic activity for hSPC1/714 and hSPC1/585 was 20.1 and 28.5%, respectively, with a 15.5- and 30.5-fold purification factor, respectively. Extrapolations suggest that this system can produce upwards of 3 mg of

crude active enzyme and hence 1 mg once purified per liter of medium. Posttranslational modifications of purified enzymes. As seen in Fig. 4C, NH 2-terminal sequencing on the 83-, 80-, and 59-kDa forms shows identical results, which are in accordance with cleavage at the expected zymogen activation site (COOH-terminal to the Arg 104Thr-Lys-Arg 107 sequence). Indeed, the deduced sequence corresponds exactly in the three cases to the mature and active hSPC1. Thus, the difference between the 83 kDa and the 80 kDa must reside either in a different level of glycosylation (see below) or in a COOH-terminal truncation of the protein chain. The hSPC1 enzyme contains three putative Asnlinked glycosylation sites. Since deficiency in oligosaccharide processing has often been observed in insect cells, we examined the glycosylation state of the purified SPC1 forms. Treatment of the hSPC1/714 83- to 80-kDa doublet with PNGase F, that removes asparagine-linked oligosaccharides, resulted in a doublet with molecular weights that shifted to 80 –76 kDa (Fig. 5A, middle panel). This same treatment on purified hSPC1/585 gives rise to a 55-kDa protein, a decrease of around 4 kDa from the original 59-kDa form, a shift similar to the one observed with the 83/80-kDa forms. This decrease in molecular weight signifies that the proteins produced with this system are glycosylated. It remains to be determined whether all three sites are N-glycosylated. We next wanted to verify the sialylated state of the molecules. As seen in Fig. 5B, no molecular weight shift was observed when SPC1/714 or SPC1/585 was treated with neuraminidase. As a control, the same conditions were used to fully desialylate commercially available human ␣ 1-antitrypsin (Fig. 5C), which

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FIG. 3. Small-scale (100 mL) purification of hSPC1/714 and hSPC1/585 from the medium of S2 cells. (A) Concentration of SPC1 activity by ammonium sulfate precipitation: enzymatic activity was measured as described under Materials and Methods using the fluorogenic substrate t-Boc-Arg-Val-Arg-Arg-MCA. (B) Western blot analysis of the proteins present in the complete culture medium and of the proteins present in the precipitate following ammonium sulfate. The proteins were first separated by SDS–PAGE and then transferred to Hybond-C membrane. Ensuing blot was analyzed by Western analysis using a specific SPC1 antibody. (C) Purification of hSPC1 forms by gel filtration chromatography. The proteins contained in the pellet from (A) were then chromatographed on a Sephacryl S-100 gel filtration column. Each fraction was tested for SPC1 activity. (D) Pooled fractions from gel filtration chromatography were further purified by FPLC on a MonoQ anion-exchange column. Enzymatic activity of each fraction is shown by the histogram bar graph. Optical density profiles at 254 nm for hSPC1/714 (continuous line) and hSPC1/585 (dotted line) are also presented along with the sodium chloride gradient (dashed line) used to elute proteins from the column.

resulted in a shift toward a lower apparent molecular weight. Kinetic analysis and stability of purified hSPC1. Heterologous expression of mammalian convertases in insect cells may result in loss of enzymatic activity or modification of kinetic parameters due to different posttranslational modifications, misfolding, or the physicochemical environment in which the protein is synthesized. To test this possibility, we performed a set of experiments to determine the Michaelis–Menten constant (K M), limiting rate (V max), turnover rate (k cat), and k cat/K M ratio of each form using fluorogenic peptide substrates. Representative plots of initial rate versus t-Boc-RVRR-MCA concentrations with hSPC1/714 and hSPC1/585 are shown in Fig. 6. As seen in Fig. 6A, crude medium from S2 cells that express hSPC1/585 or hSPC1/714 contains enzymatic activity showing a similar K M (15.2 ⫾ 0.4 and 17.7 ⫾ 0.7 ␮M, respectively). To obtain comparable V max, k cat, and k cat/V max, purified enzymes were used. Purified hSPC1/585 shows standard Michaelis–Menten kinetics with a K M of 12.0 ⫾ 1.0 ␮M

and a limiting rate of 18.0 ⫾ 0.3 nM/min of MCA produced per picomole of purified enzyme (as determined under Material and Methods) yielding a turnover rate of 0.3 s ⫺1. Purified hSPC1/714 behaved differently. At substrate concentrations over 50 ␮M, MCA production decreased, suggesting substrate inhibition. Such phenomenon was previously observed among subtilisin-like enzymes, more precisely with purified yeast kexin though not with MCA-containing substrates but with internally quenched fluorogenic substrates (20). When experimental data were fitted to an equation describing substrate inhibition (Eq. [1]), a K M of 87.9 ⫾ 7.2 ␮M and a V max of 62.5 ⫾ 4.1 nM/min per picomole of enzyme was observed. This discrepancy between the two forms prompted us to examine the possibility that at higher substrate concentrations, the enzyme tends to precipitate. Indeed, addition of 0.5 mg/mL BSA together with 0.5% Triton X-100 to the assay buffer is able to completely abolish this behavior. Although hSPC1/585 did not exhibit the same behavior as hSPC1/714, addition of BSA and detergent helped in

CHARACTERIZATION OF RECOMBINANT hSPC1 IN S2 CELLS

FIG. 4. Characterization of purified recombinant hSPC1/714 and hSPC1/585. (A) SDS–PAGE of purified hSPC1/714 (100 ng) and hSPC1/585 (1 ␮g). Protein bands were stained using GelCode Coomassie blue. (B) Western blot analysis of the same sample as in (A) using an anti-SPC1 specific antiserum as described under Materials and Methods. (C) Automated NH 2-terminal sequences of SDS– PAGE-separated hSPC1 constructs. The yield in picomoles (expressed on a logarithmic scale) of the recovered phenylthiohydantoin-amino acids at each cycle is plotted as a function of cycle number. The repetitive yield is indicated by the line following linear regression analysis and corresponds to 83.6, 95.9, and 88.4% for hSPC1/714-83 kDa (F), hSPC1/714-80 kDa (E), and hSPC1/585 (Œ), respectively. The residues occupying positions 15, 18, and 19 were not positively identified in some sequences and thus are not indicated.

reducing variations at higher substrate concentration (compare Figs. 6B and 6C). The enzyme kinetic results are summarized in Table 2. Kinetic parameters, using a second MCA substrate having the cleavage motif Arg-Thr-Lys-Arg that mimics the autocatalytic activation site of hSPC1, were also determined. This substrate has a 10-fold higher affinity toward hSPC1 compared to Boc-RVRR-MCA (see Table 2). Using this substrate, a 10-fold higher enzymatic specificity as described by the k cat/K M ratio was also obtained since k cat values were unchanged (see Table 2). Analysis of kinetic data by Hill plot revealed no cooperativity for both forms (0.99 for hSPC1/714 and 1.00 for hSPC1/585), implying that hSPC1 essentially behaves as a monomer. The kinetic parameters presented here are similar to those obtained using other expression systems (15–17) suggesting that

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hSPC1 produced by S2 cells is enzymatically equivalent to hSPC1 produced in mammalian cells. To efficiently examine the pH profile of our purified enzymes, we used two assay buffers to scan pHs between 5.8 and 10. The normalized activity measurements obtained in Hepes and Tris buffers are shown in Figs. 7A and 7B for hSPC1/714 and hSPC1/585, respectively. The normalized curves indicate that hSPC1/714 has optimal activity between 7.25 and 7.75, compared to a wider range observed for hSPC1/585 (between 6.5 and 8.0). The calcium dependence of hSPC1 forms (Fig. 7C) was determined using Hepes buffer at pH 7.5. Relatively high concentrations of calcium were required to fully activate the enzyme (0.5–2 mM) with half-maximal activation (K 0.5) at approximately 200 ␮M for hSPC1/714 and twofold lower (100 ␮M) for hSPC1/585. High concentrations of Ca 2⫹ (5–10 mM) inhibited both forms. We next wanted to characterize the stability of purified hSPC1 over time. Figure 8A is a continuous enzymatic assay of hSPC1⬘s capacity at hydrolyzing Boc-RVRR-MCA over a period of 32 h. Fitting these data to a hyperbolic curve allows us to determine the amount of substrate that could be transformed by a fixed amount of enzyme. When 1 pmol (based on weight) of both forms is incubated with 25 ␮M BocRVRR-MCA, a maximum of 12,200 ⫾ 800 and 3400 ⫾ 100 pmol of MCA can be produced by hSPC1/714 and hSPC1/585, respectively. This suggests that one molecule of hSPC1/714 cleaves, on average, 12,200 molecules of substrate before becoming inactive. It is noteworthy that hSPC1/714⬘s enzymatic activity lasted 3.6 time longer than the short form, if we assume that k cat and substrate concentration do not change significantly as the reaction proceeds. Protein stability was also assessed by incubating purified enzymes at neutral pH in buffer with calcium for 16 h at 37°C (Figure 8B). Both hSPC1/714 and hSPC1/585 show little degradation compared to control samples that were kept at ⫺20°C. It also shows that the ratio between the 83- and the 80-kDa form of hSPC1/714 is stable during the incubation and that each molecule did not have proteolytic effect on each other, as shown by coincubation of hSPC1/714 and hSPC1/585. DISCUSSION

Following transfection and antibiotic selection, S2 cells expressed recombinant hSPC1 at the milligrams per liter levels. In 1 L of crude culture medium, we estimate that the stable S2 cells produced up to 3 mg/L of unpurified, truncated hSPC1, levels that are higher than any previously used mammalian expression systems. It is higher than the CHO cells expressing SPC3 [2 mg/L (22)], recombinant vaccinia virus expressing SPC1 [⬍⬍1 mg/L (15)], and recombinant baculovirus

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structs especially with the hSPC1/714 (Fig. 2A) in this system differs from other expression systems, where it was observed that a large amount of convertase, SPC3, was retained intracellularly in baculovirus-infected Sf9 cells (4). In that sense, it more closely resembles the secretion obtained through baculovirus infection (16).

FIG. 5. Posttranslational modification of recombinant hSPC1/714 and hSPC1/585. Glycosylation state of hSPC1/714 and hSPC1/585 forms. The purified enzymes (50 –100 ng) were treated with (A) 0.5 U of PNGase F or (B) 0.5 U of neuraminidase, separated on a 8% SDS–PAGE, and transferred to nitrocellulose for Western blotting with a specific anti-SPC1 antiserum. HSPC1/714(83) refers to a FPLC fraction enriched in the 83-kDa (fraction 14, Fig. 3) form of hSPC1/714, hSPC1/714(83-80) to a fraction containing both forms (fraction 15, Fig. 3) of hSPC1/714, and hSPC1/585 to fraction 13 from Fig. 3. Commercially available serum-purified ␣ 1-antitrypsin (2 ␮g, Coomassie blue staining, Bachem Bioscience) was used as a control for the neuraminidase treatment. The protein bands displaying a decreased molecular weight due to deglycosylation or desialylation are marked with an asterisk.

expressing SPC1 [1.9 mg/L (16)]. Higher initial yields have been obtained with other convertases such as SPC2 expressed in CHO cells [6.2 mg/L (23)] or SPC3 expressed in infected Hi5 cells [6 mg/L (24)], but with a final recovery similar to the one reported here. An important feature of this system is that establishment of clonal cell lines is not necessary to achieve such level of expression. Moreover, the simple purification protocol described herein leads within 48 h to a reproducible and efficient recovery of protein amenable for structural studies. Each chromatographic step used led to a single peak of activity suggesting that S2 cells do not produce levels of proteases with trypsin-like activity that would be detected by our fluorogenic assay and/or proteases likely to degrade the expressed hSPC1. Enzymatic purity is essential if one wishes to produce enzymes for inhibitor library screening, for substrate specificity studies, or for biotechnological processes (14). Previously, it had been shown that heterologous expression of native SPC1 in various cell lines produced an intracellular doublet corresponding to the zymogen and active form of the enzyme (7,25,26). Here, with either of the constructs used, only the mature form of the enzyme is visible in cell extracts suggesting a rapid maturation of the pro-SPC1 to the active form with a slower transit to the cell surface where it is secreted. The particular efficiency of secretion of the SPC1 con-

FIG. 6. Enzymatic characterization of recombinant hSPC1/714 and hSPC1/585. Representative plots of the initial rate (V i in nM/min MCA-released) versus the amounts of fluorogenic substrate BocRVRR-MCA. The V max and K M were determined by fitting the experimental data (dark symbols) to a pseudo-first-order rate curve yielding the computed value (clear symbols) and the best fit curve. (A) Four to 20 ␮L of conditioned serum-free medium from S2-hSPC1/714 (F) or S2-hSPC1/585 (Œ) cells, respectively, were added to buffer containing various concentrations of the fluorogenic substrate and were incubated at 37°C for 1 h as described under Materials and Methods.( B) 0.4 and 0.35 pmol/assay (on a weight basis) of purified hSPC1/714 (F) and hSPC1/585 (Œ) were assayed as in (A). Data were fitted to Eq. [1] that described substrate inhibition (see text for explanation). (C) 0.6 and 0.7 pmol/assay (on a weight basis) of purified hSPC1/714 (F) and hSPC1/585 (Œ) respectively in buffer containing 0.5 mg/mL BSA and 0.5% Triton X-100.

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TABLE 2 Kinetic Parameters of hSPC1/714 and hSPC1/585

Substrate boc-RVRR-MCA

pERTKR-MCA

Enzyme hSPC1/714 hSPC1/585 hSPC1/585 hSPC1/714 hSPC1/585 hSPC1/714 hSPC1/585

a

(crude) (crude) (purified) (purified) (purified) (purified) (purified)

KM (␮M) b

V max (nM/min/pmol of enzyme)

k cat (s ⫺1)

k cat/K M ⫻ 10 4 (s ⫺1 M ⫺1)

17.7 ⫾ 0.7 15.2 ⫾ 0.4 12.0 ⫾ 1.0 21.8 ⫾ 0.8 23.0 ⫾ 0.7 2.4 ⫾ 0.2 2.3 ⫾ 0.2

n.d. n.d. 18.0 ⫾ 0.3 30.8 ⫾ 0.3 34.6 ⫾ 0.3 49.1 ⫾ 0.9 36.9 ⫾ 0.9

n.d. n.d. 0.30 ⫾ 0.01 0.51 ⫾ 0.01 0.58 ⫾ 0.01 0.82 ⫾ 0.02 0.62 ⫾ 0.02

n.d. n.d. 2.50 ⫾ 0.29 2.34 ⫾ 0.13 2.52 ⫾ 0.12 34.17 ⫾ 3.68 26.96 ⫾ 3.21

Note. n.d., not determined. a Assays containing purified enzyme include 0.5 mg/mL BSA fraction V and 0.5% Triton X-100. b Kinetic parameters were obtained by fitting experimental values to a pseudo-first-order rate curve. (See remarks concerning K cat and enzyme concentrations under Material and Methods.)

Two forms of hSPC1 were expressed in S2 cells. The first, hSPC1/714, was originally designed for efficient secretion by Molloy et al. (15) and represents the whole luminal part of the enzyme where the transmembrane domain and the cytoplasmic tail have been removed. This construction has been used extensively by different groups (15,25,26). When hSPC1/714 is expressed in S2 cells, two immunoreactive entities are secreted. The major form, 83 kDa, would be amino acids 108 through 714; the first 107 residues being the signal peptide and the proregion that is cleaved shortly after translation in the ER (7). The identity of the second form is not as clear but three elements suggested to us that it arises from a COOH-terminal truncation of hSPC1/714; (i) NH 2-terminal sequencing revealed the same amino acid sequence for both the 83- and the 80-kDa forms, (ii) treatment of the doublet with PNGase F still gave a doublet with lower apparent molecular weight, and (iii) overexpression of native SPC1 in mammalian cells (26,28 –30) and in S2 cells (unpublished results) has generated a shed form of the enzyme. The same mechanism may be responsible for this lower molecular form; hSPC1/714 may be cleaved at the C-terminus by a protease during its biosynthesis en route to the cell surface. Furthermore, we show herein that the incubation of hSPC1/714 and hSPC1/585 for prolonged periods of time in conditions favorable to hSPC1 activity does not lead to an intra- or intermolecular cleavage of SPC1. Moreover, inspection of the primary sequence of mature SPC1 reveals no SPC1 recognition site for cleavage but only single arginine residues at positions 693, 701, and 703 are found. They do not constitute the minimal Arg-X-X-Arg recognition motif though furin was shown to be able to cleave albeit much less efficiently at single Arg residues using small synthetic substrates (21). These results support previous finding that, in mammalian cells and possibly in insect cells, SPC1 shedding is sensitive to acidotropic agents indicating that this event takes place in an acidic compart-

ment of the cells (31). The second form, hSPC1/585, has its cysteine-rich region been removed and is thus closer to the minimal active form previously reported for SPC1 that is equivalent to hSPC1/577 (32). When expressed in S2 cells, it is secreted as a single glycosylated 59-kDa protein. The absence of heterogeneity makes it amenable to initiate crystallization studies. Furthermore, it has the same kinetic parameters of hSPC1/714. The anomalous behavior of purified hSPC1/714 at higher substrate concentration may indicate that hSPC1/714 once purified is either unstable in the absence of other proteins present, for example, in the crude sample or through addition of albumin, and/or is prone to aggregation or precipitation. In both cases, this would lead to a loss of active enzyme, hence the observed decline in hydrolysis rates. Though either explanation does not rule out substrate inhibition, a recent report by Krysan and co-workers (33) clearly demonstrates a substrate inhibition behavior with hexapeptide, but not with tetrapeptide substrates, like the one used in this study. We have shown an easy and efficient manner in which to circumvent the problem as addition of BSA and detergent restores pseudo-firstorder kinetics. Previous studies on the posttranslational modifications of bovine SPC1 in MDCK cells have revealed that it is synthesized as a glycosylated protein with terminal sialylated carbohydrate moieties (31). Since many studies on dipteran, orthopteran, and lepidopteran cells including Sf9 cells (34) have led to the general hypothesis that insect cells do process oligosaccharides to complex sugars but that sialylation is deficient in these cells, we have tested the glycosylation/sialylation state of our purified proteins. The shift in molecular weight of purified hSPC1 forms following PNGase treatment clearly indicates the N-glycosylated nature of the produced proteins. Although it would appear that these glycoproteins would not be sialylated as shown by the insensitivity to neuraminidase, further

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the CRR and then influence kinetic parameters remains valid. Second, the results shown herein demonstrate that deletion of this domain renders the enzyme less sensitive to regulation by pH and calcium giving rise to a broader optimum pH range and lower requirement for calcium. A study by Zhou and co-workers (9) has shown that the P-domain regulates the stability, calcium dependence, and pH dependence of the convertases SPC2 and SPC3. It is conceivable that the P-domain of all convertases plays this role and that the CRR of SPC1 and the other SPCs containing a CRR complements this regulation. A tight control of SPC1 activity by these factors may be important. Indeed, Anderson and co-workers (5) clearly show that pH and calcium regulate SPC1 maturation from the ER to the TGN. They demonstrated that, in the neutral environment of the ER, the cleaved propeptide remains bound to the protease and functions as an autoinhibitor. Exposure of the SPC1–propeptide complex to a mildly acidic and free calcium-containing environment, characteristic of the TGN, results in a second cleavage within the propeptide releasing the fragments and the fully active enzyme. As shown herein, it is also clear that the CRR of SPC1 keeps the enzyme active for a longer period of time. The CRR may thus act as a

FIG. 7. Determination of pH optimum and calcium dependence of purified hSPC1/714 and hSPC1/585. The activity of (A) hSPC1/714 and (B) hSPC1/585 over a broad pH range was determined using a fluorometric assay with Boc-RVRR-MCA (see Materials and Methods). Assays were conducted as described, except that 100 mM Hepes (E) and 100 mM Tris base (Œ) were used as buffers at pH values indicated. The activity measurements for each buffer were normalized to the maximum value obtained in that buffer (⫽100% of maximum activity). (C) Dependence of hSPC1/714 (F) and hSPC1/585 (Œ) activity as a function of calcium concentration was determined using the indicated concentrations of CaCl 2 added to the reaction mixture. The data were normalized to the maximum activity observed at 1 mM calcium. Each point represents the average of two separate determinations performed in duplicate.

experimentation, such as lectin blotting, to detect the presence of sialic acid would be required. Functional analysis of the purified forms has shown two major aspects. First, both forms are undistinguishable from each other from an enzymatic point of view. In fact, truncation on the CRR had no effect on the affinity and efficiency of the enzyme to cleave small fluorogenic peptides. However, the possibility that larger substrates like precursor proteins interact with

FIG. 8. Stability of purified hSPC1 enzyme forms. (A) Stability of enzymatic activity over time. One picomole of purified enzyme was incubated in assay buffer in the presence of 25 ␮M Boc-RVRR-MCA in 1.0 mL at 37°C. Activity was measured at various times after initiation of the assay. Experimental data were fitted to an hyperbolic curve yielding the computed values (clear symbols) and the best fit curve. A maximum MCA production of 12.2 ⫾ 0.8 and 3.4 ⫾ 0.1 nmol/pmol of enzyme for hSPC1/714 (F) and hSPC1/585 (Œ) were obtained, respectively. (B) To assess protein stability, 100 ng of hSPC1/714 and hSPC1/585 was either kept at ⫺20 oC or incubated at 37°C for 16 h in assay buffer. Samples were analyzed by Western blot.

CHARACTERIZATION OF RECOMBINANT hSPC1 IN S2 CELLS

stabilizing domain in addition to the P-domain. These results together with the presence of CRR in many proteins, including three members of the convertases family, suggest that this domain plays an important role in these convertases. For example, the CRR of SPC4, which has five repeated cysteine motifs, has been associated with retention within the neuroendocrine cell line AtT-20, but not in the fibroblastic cell line hEK-293 (35) implying the cell-specific role of this domain. The CRR is similar to the one found in the TNF␣/EGF family of receptors and to a candidate Marfan syndrome allele (11). New insight into the role of the CRR in these proteins may be useful in delineating the importance of this domain in SPC1. Recently, a study revealed that mutated cysteine residues involved in disulfide bonds in the CRR of TNF receptor (p55) causes a defective activation-induced cleavage of the receptor that leads to dominantly inherited syndrome of episodic fever and inflammation (36). It may suggest that modification of the structural organization of the CRR could result in abnormal biosynthesis of various SPC forms (e.g., shedding may be affected by modification of the CRR). All expression systems previously described suffer from a certain number of disadvantages. Both the vaccinia virus- and baculovirus-based systems are laborious in preparation of the recombinant virus and expression/purification is sometimes difficult. The CHO system needs many weeks of selection–amplification and is expensive and time consuming. Finally, bacteria- or yeast-based expression systems have failed to produce active convertases even if a counterpart of the mammalian convertases exists in yeast (kexin). The expression system used herein for the high-level expression of recombinant hSPC1 has been demonstrated to be simple, quick, and effective and able to produce milligrams of active enzyme. The purified enzymes show comparable kinetic parameters to the ones obtained with other mammalian expression systems, similar calcium and pH dependencies, and posttranslational carbohydrate modifications. This system is presently used with success to express and purify other members of the subtilisin proprotein convertases family and may be a valuable tool for the understanding and characterization of these important enzymes. ACKNOWLEDGMENTS This study was supported by Research Grants MT-14194 and MT-15550 (R.D.), MT-14766 (C.L.,) and MT-13755 (R.L.) from the Medical Research Council of Canada and by Grant 0GP0171374 (R.L.) from the Natural Sciences and Engineering Research Council of Canada. J.-B.D. is supported by the Fonds de la recherche en sante´ du Que´bec (FRSQ). R.D. and R.L. are both scholars of the FRSQ. We wish to thank Karin Losito for excellent technical assistance.

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