Mol. Hum. Reprod. Advance Access originally published online on December 17, 2008
Molecular Human Reproduction 2009 15(2):105-114; doi:10.1093/molehr/gan077
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Recombinant expression and affinity purification of a novel epididymal human sperm-binding protein, BSPH1
1Research Centre, Maisonneuve-Rosemont Hospital, University of Montreal, 5415 Boulevard de L’Assomption, Montreal, Quebec, Canada H1T 2M4 2Department of Biochemistry, University of Montreal, Montreal, Quebec, Canada H1T 2M4 3Department of Medicine, University of Montreal, Montreal, Quebec, Canada H1T 2M4
4 Correspondence address. Tel: +1-514-252-3562 or +1-514-252-3400 (ext. 3329); Fax: +1-514-252-3430; E-mail: puttaswamy.manjunath{at}umontreal.ca
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Abstract |
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Mammalian sperm undergo a series of maturation steps before acquiring fertilization competence. Our previous work demonstrated the importance of binder of sperm (BSP) proteins in bovine sperm capacitation. Recent studies identified a BSP-homologous DNA sequence in the human genome (BSPH1) and mRNA expression in the epididymis. The aim of this study was to develop an efficient method to express and purify recombinant human BSPH1. BSPH1 accumulates in inclusion bodies when expressed with an N-terminal hexahistidine tag in BL21 (DE3) Escherichia coli cells. Similar to other BSP proteins, BSPH1 contains two fibronectin type-II (Fn2) domains, each consisting of two disulfide bonds. Therefore, when expressed in Origami B (DE3)pLysS cells, a strain favouring disulfide bond formation, an improvement in soluble protein yield was observed. However, protein was aggregated, which complicated subsequent purification steps. Expression of glutathione-S-transferase-tagged BSPH1 in both cell types also led to accumulation in inclusion bodies. Finally, successful production of soluble and active protein was achieved when BSPH1 was expressed as a His6-thioredoxin-tagged protein. Recombinant protein bound phosphatidylcholine liposomes, low-density lipoproteins and human sperm, therefore displayed binding activities common to all BSP-family proteins, which may indicate similar biological function(s). This approach was also successful in producing the murine orthologue of BSPH1 in the soluble and active form. Thus, fusion to thioredoxin and expression in Origami B (DE3)pLysS cells may constitute a strategy applicable to all BSP-family proteins, and possibly to other proteins containing Fn2 domains. This work is important to elucidate the role of BSPH1 in human sperm functions and fertility.
Key words: epididymal protein/binder of sperm (BSP) proteins/fibronectin (Fn)2 domains/recombinant protein expression/thioredoxin
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Introduction |
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Sperm cells, carriers of the paternal contribution to the future embryo’s genome, are transcriptionally and translationally silent cells; a consequence of their highly condensed chromatin and the presence of little cytoplasm. Therefore, modifications inflicted upon sperm during their transit through the epididymis and female reproductive tract are brought about by secreted proteins, which associate with the sperm plasma membrane. In order to fertilize an oocyte, mammalian sperm must go through two successive maturation steps; the first occurring in the epididymis, referred to as epididymal maturation, and the second taking place inside the female reproductive tract, called capacitation. The bulk of modifications undergone by sperm during epididymal maturation affect the plasma membrane, notably with the addition and rearrangement of surface proteins necessary for sperm to undergo capacitation and interact with the oocyte (Cooper, 1995; Jones, 1998).
Following epididymal maturation and ejaculation, mammalian sperm undergo a complex series of modifications inside the female reproductive tract, collectively referred to as sperm capacitation (Austin, 1951; Chang, 1951). Events occurring during capacitation include changes in the plasma membrane lipid composition, increased permeability to Ca2+, increased intracellular pH, redistribution of surface proteins, increased sperm motility, increased adenylyl cyclase activity and cyclic AMP as well as an increased tyrosine phosphorylation of a group of signalling proteins (for reviews, see de Lamirande et al., 1997; Visconti and Kopf, 1998; Visconti et al., 2002). A family of sperm-binding proteins secreted by the seminal vesicles has been shown to be essential for capacitation in the bovine species (Manjunath and Therien, 2002). These proteins, originally named bovine seminal plasma proteins, bind to sperm upon ejaculation due to their interaction with sperm membrane choline phospholipids (Desnoyers and Manjunath, 1992; Manjunath et al., 1994). Very recently, the genes encoding BSP proteins were renamed, the BSP acronym now standing for ‘binder of sperm’ (Manjunath et al., 2008). When sperm arrive at the site of fertilization, BSP proteins interact with follicular and oviductal fluid capacitation factors, high-density lipoproteins (HDL) and glycosaminoglycans, and induce cholesterol efflux and intracellular signalling pathways, respectively, significantly contributing to the capacitated state of spermatozoa (reviewed in Manjunath and Therien, 2002; Manjunath et al., 2007).
BSP proteins have been thoroughly characterized at the biochemical, structural and molecular levels (Kemme and Scheit, 1988; Manjunath et al., 1988; Salois et al., 1999). The signature of the BSP family of proteins is the presence of two tandemly arranged fibronectin type-II domains (Fn2 domains), similar to those found in the gelatin-binding domain of fibronectin (Fan et al., 2006). These domains each contain four cysteine residues, which in turn form two disulfide bridges per domain (Fig. 1). Thus, each BSP protein contains four disulfide bonds. Fn2 domains are responsible for the binding of BSP proteins to HDL (Manjunath et al., 1989), glycosaminoglycans (Therien et al., 2005), choline phospholipids (Desnoyers and Manjunath, 1992), collagen and gelatin (Manjunath et al., 1987). BSP-homologous proteins are also present in the seminal fluids of boar, stallion, goat, ram and bison (reviewed in Manjunath et al., 2007). In all the previously mentioned species, BSP-like proteins are present in the seminal fluid in relatively large quantities (ranging from 1% to 60% of total seminal plasma proteins). Consequently, the study of their biological functions was made possible by affinity purification of the native proteins directly from seminal plasma, followed by subsequent incubation of the purified proteins with sperm for functional assays (Therien et al., 1995, 1997; Lusignan et al., 2007). Recently, we have shown that BSP homologues are also expressed in mice and human (Lefebvre et al., 2007). However, in these species, BSP-like proteins are expressed in the epididymis and represent a minute proportion of seminal plasma proteins. This renders the task of purifying sufficient quantities of the native protein quite complex, thus favouring the use of recombinant technology.
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DNA sequences containing one or more repeats of Fn2 domains are found in more than 100 genes in numerous mammalian genomes (Fan et al., 2006). The phylogeny of Fn2 domain-containing proteins has been thoroughly described (Fan et al., 2006), and indicates the existence of at least 12 different families encompassing more than 50 proteins. Despite the presence of Fn2 domains in a wide array of proteins, such as extracellular matrix proteins, membrane-associated proteins, matrix metalloproteases (MMPs), seminal fluid proteins and serine proteases (Ozhogina et al., 2001), very few of these have been successfully expressed in a recombinant system. Moreover, of those expressed, very few studies report high-yield soluble expression, and most resorted to the solubilization and refolding of recombinant protein from insoluble inclusion bodies (Banyai et al., 1990; Tordai and Patthy, 1999; Jani et al., 2005). Hence, efforts aimed at developing an efficient strategy to produce soluble recombinant Fn2 domain-containing proteins are warranted.
The aim of the present study was to express and purify the recombinant human BSP protein, BSPH1 (previously named hBSPH1), which contains two Fn2 domains, to facilitate the examination of its biological role in sperm functions. Since BSP proteins are essential for bovine and porcine sperm capacitation, it is justified to hypothesize that the human counterpart would be involved in human sperm functions. However, the unavailability of native protein has made the confirmation of this hypothesis impossible. The soluble expression of recombinant human BSPH1 and murine Bsph1 (orthologue of BSPH1) will allow the investigation of the role of this protein in sperm functions and fertility. In addition, BSP proteins have been shown to be detrimental in the context of sperm storage and cryopreservation (Manjunath et al., 2007). Therefore, purified recombinant proteins will be useful in investigating the effect of BSPH1 and Bsph1 on sperm storage in the mouse and human species, respectively.
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Materials and Methods |
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Materials
Plasmids pET15b and pET32a, as well as Escherichia coli host strains BL21 (DE3)pLysS and Origami B (DE3)pLysS and His-bind resin were from Novagen (EMD Biosciences, La Jolla, CA, USA). pGEX-5x-1 expression vector, Sephacryl S-100 and Taq DNA polymerase were from GE Healthcare (Baie d’Urfé, QC, Canada), while Pfu DNA polymerase was from Fermentas (Burlington, ON, Canada). Restriction enzymes were from New England BioLabs (Beverly, MA, USA). QIAprep Spin Miniprep Kit and Qiaex II gel extraction kit were from Qiagen (Mississauga, ON, Canada). TA Cloning kit and T4 DNA ligase were from Invitrogen (Carlsbad, CA, USA). For western blotting, the His-Probe monoclonal antibody was from Santa Cruz (Santa Cruz, CA, USA), whereas the goat anti-mouse IgG and goat anti-rabbit IgG were from Bio-Rad (Mississauga, ON, Canada), and the chemiluminescence reagent was from Perkin–Elmer (Boston, MA, USA). Freund’s adjuvants (complete and incomplete) were purchased from Sigma–Aldrich (Oakville, ON, Canada). Complete Mini, EDTA-free protease inhibitor tablets were from Roche (Manheim, Germany). Finally, the B-PER Bacterial Protein Extraction Reagent was from Pierce (Rockford, IL, USA), and all other chemicals used were of analytical grade and obtained from commercial suppliers.
Cloning of cDNA sequences into expression vectors
For expression of BSPH1 with an N-terminal His-tag, human epididymal cDNA was used as a template for polymerase chain reaction (PCR) amplification of BSPH1. In order to clone into the expression vector pET15b, the following oligonucleotide primers were employed: BSPH1-F(Nde1) 5'-GCG CAT ATG TGC ATC TTC CCT GTT ATT TTA AAT G-3' and BSPH1-R(BamH1) 5'-GCA GGA TCC TCA TTC ACA GTA TTT CCA AAT TCG-3'.
For expression of BSPH1 with an N-terminal glutathione-S-transferase (GST)-tag, human epididymal cDNA was once again used as a template for PCR, using the following primers: BSPH1-F(BamH1) 5'-GCG GGG ATC CTC ATC TTC CCT GTT ATT TTA AAT G-3' and BSPH1-R(Xho1) 5'-CGC CTC GAG TCA TTC ACA GTA TTT CCA AAT TCG-3'. The restriction sites added during PCR amplification allowed subsequent cloning into to pGEX-5x-1 expression vector.
For expression of BSPH1 fused to thioredoxin, the pET15b-BSPH1 plasmid was used as a DNA template. The following primers were used, which allowed subsequent cloning into the expression vector pET32a: BSPH1-F2 (BamH1) 5'-GCA GGA TCC ATC TTC CCT GTT ATT TTA AAT G-3' and BSPH1-R(Xho1).
PCR amplification was performed using Pfu DNA polymerase, under the following conditions: 94°C, 3 min; 33 cycles of 94°C, 45 s; 60°C, 45 s; 72°C, 1 min and one cycle of 72°C, 7 min. Taq DNA polymerase was added to PCRs during the final elongation step, to allow subsequent TA sub-cloning into pCR2.1. Sub-cloned BSPH1 sequences were excised from pCR2.1 using the appropriate restriction enzyme pairs mentioned above, while the expression vectors were linearized using the same enzymes. Digested inserts and vectors were run on a 1% agarose gel, gel-purified using the Qiaex II gel extraction kit and then ligated overnight using T4 DNA ligase. Ligation reactions were transformed into competent DH5
cells, and plasmid DNA was isolated using the QIAprep Spin Miniprep Kit. Sequences were confirmed using the Big Dye® Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, CA, USA) and the ABI PRISM® 3100 Genetic Analyzer (Applied Biosystems).
Protein expression in E. coli
Plasmids were transformed into BL21 (DE3)pLysS or Origami B (DE3)pLysS competent cells using standard methods (Sambrook and Russel, 2001). Transformed bacteria were grown overnight on LB-agar plates containing 100 µg/ml ampicillin, after which single colonies were used to inoculate liquid LB medium containing the same antibiotic. For protein expression, 300 ml to 1 l liquid LB medium was inoculated with 1/100 volume of overnight culture; bacteria were grown at 37°C with shaking at 200 rpm until the OD600 reached 0.6–0.8 and then the lactose analogue, isopropyl-β-D-thiogalactopyranoside (IPTG) was added to induce expression. An IPTG concentration of 1 mM was used to induce expression of His-tagged and thioredoxin-fused BSPH1 in BL21 (DE3)pLysS cells for 3 h at 30°C, as well as in Origami B (DE3)pLysS cells when induction was performed at 16°C for 16 h. In the case of 3–4 h inductions of Origami B (DE3)pLysS cells at 30°C, 0.2 mM IPTG was used, whereas 0.1 mM IPTG was used to induce expression of GST-tagged BSPH1 in both cell types. Following induction, cells were harvested by centrifugation at 5000 xg for 10 min at 4°C. One milliliter of aliquots of bacterial suspensions were removed before and after induction, pelleted by centrifugation and resuspended in 100 µl sample buffer. Ten microliter of each was used in electrophoresis and western blot experiments.
Nickel affinity chromatography
Cell pellets were resuspended in B-PER Bacterial Protein Extraction Reagent and processed according to manufacturer’s instructions, except that the cell lysate was subjected to sonication (three cycles of 10 s on ice) before centrifugation at 15 000 xg for 15 min to separate the soluble and insoluble fractions. Supernatant was loaded onto a His-Bind column, prepared and equilibrated according to manufacturer’s instructions. Unless otherwise specified, washing and elution were performed as described in the Novagen protocol handbook. All samples were analysed by SDS–PAGE and western blotting.
Protein electrophoresis and western blotting
SDS–PAGE in 15% gels was performed according to Laemmli (1970) using the Mini Protean 3 apparatus from Bio-Rad. Gels were either stained with Coomassie Brilliant Blue, or transferred electrophoretically to Immobilon-P PVDF membranes (Millipore, Nepean, ON, Canada). Immunodetection was performed using either a His-Probe monoclonal antibody at a concentration of 1:1000, affinity-purified antibodies against a synthetic peptide corresponding to the 15 C-terminal amino acids of the deduced sequence of BSPH1 (1:1000), or affinity-purified antibodies against His6-tagged recombinant BSPH1 (1:2000). Goat anti-mouse IgG (1:3000) and goat anti-rabbit IgG (1:10 000) were used as secondary antibodies, then blots were revealed using a chemiluminescence reagent and a Fuji LAS-3000 image analyzer (Fujifilm; Stamford, CT, USA).
Generation of antiserum against human BSPH1
A 15 amino acid peptide corresponding to the C-terminus of BSPH1 (SLTKNFNKDRIWKYCE) was synthesized and conjugated to keyhole limpet hemocyanin at the Sheldon Biotechnology Centre (Montreal, QC, Canada). New Zealand rabbits were injected hypodermically with a mixture of 200 µg conjugated peptide dissolved in 100 µl 50 mM PBS, 400 µl sterile 0.9% NaCl and 500 µl Freund’s complete adjuvant. Boosts were performed at 20-day intervals over a period of 88 days, with 200 µg of the same antigen in the same mixture, except using Freund’s incomplete adjuvant. Bleedings were performed 15 days after each injection. Antiserum from the third boost was used for the present study.
In the case of antibodies raised against the whole protein, His6-tagged BSPH1 was expressed as described earlier and inclusion bodies were isolated according to the instructions in the Novagen protocol handbook. For each injection, 
175 µg protein equivalent was run on a 10% SDS–PAGE gel. Fifteen microgram of protein was run in another lane, which was cut from the gel and stained with Coomassie Brilliant Blue. After staining, the gels were aligned and the portion of the unstained gel corresponding to BSPH1 protein was excised. Gel slices were minced in 1 ml saline solution and homogenized using a polytron. An equal volume of Freund’s complete (first injection) or incomplete (subsequent injections) adjuvant was added and the solution was emulsified. New Zealand rabbits were immunized as described earlier. Both antisera were passed through a Protein-A-Sepharose column to purify the antibodies. Using the concentrations mentioned earlier, both antibodies gave similar results in western blots. Animals were treated in accordance with the guidelines of the Canadian Council of Animal Care.
Binding to phosphatidylcholine liposomes
Liposomes of phosphatidylcholine (PC) were prepared as described in (Desnoyers and Manjunath, 1992). Briefly, 8 mg PC (Doosan Serdary Research Laboratories; Englewood Cliffs, NJ, USA) in chloroform was evaporated under N2 until a thin film was formed in the bottom of a glass test tube (30 min, 10–20 psi). PC was then resuspended in 2 ml Buffer A (10 mM Tris–HCl, 100 mM KCl, pH 7.5) and the tube was sonicated in a Branson Ultrasonic water bath (Model 3510) for 20 s at room temperature (lipids formed an opalescent suspension). Large multilamellar liposomes were sedimented by ultracentrifugation at 100 000 xg (32 300 rpm), at 25°C for 30 min in a Sorvall T-865 rotor. The pellet containing liposomes was resuspended in Buffer B (10 mM Tris–HCl, 100 mM KCl, 2.5 mM MgCl2, pH 7.5), to obtain a lipid concentration of 4 mg/ml. To verify the binding of BSPH1 (20 µg), BSA (10 µg) and crude BSP proteins (cBSP; 20 µg) to PC liposomes, the indicated amounts of protein were incubated with liposomes (equivalent to 300 µg PC), in a total volume of 300 µl Buffer B. Incubations were carried out for 30 min at room temperature, after which liposomes were sedimented once again at 100 000 xg (32 300 rpm), at 25°C for 45 min. Equivalent fractions of supernatant (proteins precipitated with trichloroacteic acid) and pellet were analysed by SDS–PAGE.
Binding to egg yolk low-density lipoproteins (LDL)
LDL were isolated from hen’s egg yolk as described previously (Manjunath et al., 2002), dialysed extensively against 10 mM Tris–HCl, pH 7.4 and dosed according to the modified Lowry procedure (Markwell et al., 1978). Purified Trx-BSPH1 (150 µg), Trx (150 µg) or crude BSP proteins (100 µg) were incubated with 200 µg LDL for 90 min at room temperature on a nutating platform. The density of the protein–lipoprotein solution was then increased to 1.21 g/l using KBr and the solutions were transferred to Quick Seal tubes and ultracentrifuged for 18 h at 366 912xg, 20°C. After centrifugation, the samples were separated into top, middle and bottom fractions and analyzed by western blotting with the appropriate antibodies.
Binding to human sperm
Human ejaculated sperm were obtained from healthy volunteers, allowed to liquefy at 37°C for 30 min and sperm were counted. Sperm were separated from seminal plasma by centrifugation at 1500 xg, 10 min, RT and washed three times with 1 ml PBS. Purified Trx-BSPH1 or Trx alone (150 µg) were incubated with 150 x 106 washed human sperm for 1 h at RT on a nutating platform. As a control, sperm were incubated in PBS in the absence of protein. The sperm suspensions were then centrifuged 1500 xg, 10 min, RT and the supernatant was removed and kept aside for analysis. Sperm were then washed twice with 1 ml PBS, then incubated with 200 µl lysis buffer (25 mM Tris–HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA, 2% SDS, 1% Triton-X-100, 50 mM DTT, protease inhibitor cocktail) and sonicated for 10 s at 50% amplitude. Lysed sperm were then centrifuged at 10 000 xg, 15 min, 4°C, in order to separate the supernatant containing the sperm-extracted proteins from the pellet containing cell debris. Hundred microliter Laemmli buffer was added to the pellet, which was then resuspended and boiled. All fractions, including 20 µl of the pellet fraction was loaded onto the gel and analysed in a western blot with the appropriate antibodies.
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Results and discussion |
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The production of soluble and highly purified recombinant eukaryotic proteins in E. coli can be a difficult task, due to numerous obstacles including inclusion body formation, misfolding and protein aggregation, which can mainly be attributed to the absence of eukaryotic chaperones. Nevertheless, E. coli remains the expression host of choice due to its rapid growth, high-level protein production, ease in manipulation, as well as low cost. The most commonly used method for protein expression in E. coli is to produce the recombinant protein in fusion with an affinity tag, rendering subsequent affinity purification a reasonably simple task (for a review, see Terpe, 2003). The polyhistidine (His)-tag is frequently used due to its small size (6–10 amino acid residues), which rarely interferes with protein activity, thus precluding the need for tag removal, as well as the possibility of a one-step purification method using immobilized metal-affinity chromatography (Porath et al., 1975). However, many proteins are produced as insoluble aggregates, which can sometimes be overcome by fusion to soluble protein partners, such as gluthathione-S-transferase), maltose-binding protein, thioredoxin and many others (for a review, see Terpe, 2003). Success in producing soluble, active protein is largely target protein-specific; therefore, there is no magical protocol applicable to every protein. In the absence of previously published protocols, production of soluble recombinant proteins becomes somewhat of a ‘trial and error’ exercise.
Expression of His6-tagged human BSPH1 in BL21 (DE3)pLysS E. coli
Our initial strategy to produce recombinant BSPH1 was to express it in fusion with an N-terminal hexahistidine tag in BL21 (DE3)pLysS E. coli cells, the most common method to produce recombinant proteins. However, as seen in Fig. 2A, the majority of His-tagged BSPH1 expressed from the pET15b expression vector accumulated in inclusion bodies. This is most probably due to the presence of four disulfide bridges in the predicted protein structure (Fig. 1). The E. coli cytoplasm is not an ideal environment for the oxidation of sulfhydryl groups to form stable disulfide bonds. Consequently, proteins harbouring multiple disulfide bonds are often incorrectly folded when expressed in E. coli. In fact, previous attempts at expressing Fn2 domain-containing proteins in E. coli also resulted in the accumulation of these proteins in inclusion bodies (Banyai et al., 1990; Tordai and Patthy, 1999; Jani et al., 2005). Protein expression in the form of inclusion bodies can offer certain advantages. Primarily, recombinant protein trapped in inclusion bodies can accumulate to levels representing more than 30% of the cellular proteins. They can easily be separated from other organelles by low-speed centrifugation, protect proteins from proteolytic degradation and contain very few contaminating proteins (high purity) (Singh and Panda, 2005). However, in order to recover active protein, the inclusion bodies must first be solubilized and the recombinant protein correctly refolded and purified. Optimizing refolding conditions can prove to be a very time-consuming and tedious task. Nevertheless, we attempted a refolding strategy for His-tagged BSPH1, since the protein was abundant, and refolding of recombinant Fn2 domain-containing proteins, MMP-2 and fibronectin, has already been accomplished (Banyai et al., 1990; Tordai and Patthy, 1999; Jani et al., 2005). Incubation with reducing agents, such as 5 mM dithiothreitol (DTT) or 10 mM tris(hydroxypropyl)phosphine, in addition to 8 M urea, was necessary to promote solubilization of the inclusion bodies (data not shown). Dialysing denatured and reduced protein against buffer devoid of urea resulted in considerable protein precipitation and dialysis against buffer containing 2.5% or 5% glycerol kept some protein in solution, although most of it precipitated (data not shown). Examples in the literature show that disulfide-containing proteins require a more complex refolding strategy to allow proper formation of disulfide bonds (Fischer et al., 1992; Vallejo and Rinas, 2004; Leung and Ho, 2006). In our hands, the approach that achieved the most success in obtaining soluble protein was a step-wise dialysis, which allowed the gradual removal of urea and included a redox system (mixture of oxidized and reduced glutathione) at an intermediate urea concentration (2 M). In addition, a low protein concentration (100 µg/ml) was used to minimize aggregation and promote refolding. Although a significant quantity of recombinant His-BSPH1 was rendered soluble using this method, the ‘refolded’ protein did not exhibit properties characteristic of BSP-family proteins, such as binding to gelatin, phosphorylcholine and sperm (data not shown), suggesting that it had not regained its native conformation. This is not uncommon, as protein solubility is not an absolute indicator of correct folding (Dyson et al., 2004; Vallejo and Rinas, 2004). After refolding from inclusion bodies, some recombinant proteins show no activity despite being in the soluble form (Huang et al., 1998).
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Expression of His6-tagged human BSPH1 in Origami B (DE3)pLysS E. coli
In a second attempt to produce soluble His-BSPH1, we expressed pET15b-BSPH1 in the Origami B (DE3)pLysS bacterial strain from Novagen. In contrast to most E. coli strains, this genetically modified strain (
trxB and gor) favours the formation of disulfide bonds in its oxidizing cytoplasm (Prinz et al., 1997; Bessette et al., 1999). When cells were induced at 30°C, recombinant protein accumulated in inclusion bodies (Fig. 2B), as seen with BL21 (DE3)pLysS cells. However, when induction was performed at 16°C for 16 h, a significant increase of soluble His-BSPH1 was observed (Fig. 2C). The Tenebrio molitor thermal hysteresis protein, which contains 16 cysteines, was successfully expressed in this strain at low induction temperature and extended induction times (Bar et al., 2006). The soluble fraction obtained after cell lysis was subjected to nickel affinity chromatography, which allowed the recovery of partially purified protein (data not shown). Due to the presence of several contaminants in the His-Bind eluate, we sought to further purify the recombinant protein by reversed-phase high-performance liquid chromatography (rp-HPLC). However, His-tagged BSPH1 eluted from the HPLC column in multiple peaks (data not shown). This suggested that the protein was aggregated, and that aggregates of different sizes eluted in different peaks due to their different hydrophobicities. It has been proposed that the presence of imidazole often results in protein aggregation (Hefti et al., 2001). Thus, we resolved to attempt purification of His-BSPH1 directly from cell lysates using gel-filtration chromatography, in order to separate our target protein according to its molecular weight. Whether we separated lysate proteins under basic (50 mM ammonium bicarbonate) or acidic (50 mM acetic acid) conditions, our protein consistently behaved like a high molecular weight aggregate, eluting in the first peak (data not shown). Thus, it seemed that His-BSPH1 was incorrectly folded, leading to protein aggregation, possibly due to hydrophobic interactions or intermolecular disulfide bonding.
Expression of GST-tagged human BSPH1
The 26-kDa GST-tag is also widely used as a fusion partner in the production of recombinant proteins in E. coli. This large tag confers several advantages, such as the possibility of single-step purification using immobilized glutathione (Smith and Johnson, 1988), as well as enhanced solubility and protection against intracellular protease cleavage. We cloned the cDNA encoding BSPH1 into the pGEX-5x-1 expression vector, which adds a N-terminal GST-tag, and induced expression of the fusion protein in BL21 (DE3)pLysS and Origami B (DE3)pLysS cells with IPTG. The GST-BSPH1 molecule was completely insoluble in BL21 (DE3)pLysS (data not shown) and in Origami B (DE3) pLysS (Fig. 2D).
Expression of His6-thioredoxin-tagged human BSPH1
Fusion of recombinant proteins to E. coli thioredoxin (trxA) was first undertaken with the rationale that it was a protein normally present in the E. coli cytoplasm, and that it could be overexpressed in the soluble form to levels nearing 40% of cellular proteins (Lunn et al., 1984; LaVallie et al., 1993). High-level expression of a soluble and functional Fn2 domain of MMP-2 was achieved in E. coli, using fusion to thioredoxin (Peisley and Gooley, 2007), validating an attempt at expressing BSPH1 as a thioredoxin fusion. When expressed in BL21 (DE3)pLysS cells at 30°C (Fig. 3A) or at room temperature (data not shown), the thioredoxin-BSPH1 fusion protein was mostly insoluble (>90%), although part of it could be seen in the soluble fraction. A considerable increase in soluble protein (70% of expressed rBSPH1 found in the supernatant) was obtained when expression was induced in Origami B (DE3)pLysS cells, irrespective of whether induction was performed for 4 h at 30°C or for 16 h at 16°C (Fig. 3B). The absence of thioredoxin reductase in the genetically modified strain allows for the accumulation of oxidized thioredoxins and the formation of disulfide bridges in the bacterial cytoplasm (Prinz et al., 1997; Stewart et al., 1998; Bessette et al., 1999). As seen in Fig. 3B, a significant amount of recombinant protein remained insoluble in inclusion bodies. Soluble yield could potentially be improved by co-expression of human protein disulfide isomerase (PDI), which would catalyse disulfide bond formation and isomerization. Zhao et al. (2000) expressed the human cardiac-specific homeobox protein, which contains six cysteine residues, in E. coli, and obtained 50% soluble protein. When co-expressed with PDI, soluble yield increased to >90%. Since our method allowed the recovery of sufficient soluble protein, we did not seek to further enhance soluble expression. The pET32a expression vector used to express BSPH1 as a thioredoxin fusion also adds a hexahistidine tag; therefore, we could use Nickel-affinity chromatography for purification (Fig. 3C). As seen in previous attempts at purifying recombinant BSPH1 using elution with 1 M imidazole, eluted proteins were prone to aggregation and precipitation during storage (not shown). However, we found that elution with a lower imidazole concentration (400 mM) was sufficient to recover our protein from the column, and protein remained in solution during extended storage periods (more than 2 weeks) at 4°C.
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Expression of His6-thioredoxin-tagged murine Bsph1
We also sought to determine whether the strategy employed to produce soluble recombinant BSPH1 could be extrapolated to other BSP-family proteins. The same procedure was repeated to express the murine orthologue of BSPH1, Bsph1 (previously named mBSPH1). As seen in Fig. 3D, murine Bsph1 was also produced in the soluble form in significant quantities when fused to thioredoxin.
Binding properties of recombinant human BSPH1
Binding to phosphatidylcholine liposomes
In order to verify if recombinant BSPH1 was active, its ability to bind phosphatidylcholine (PC) liposomes was verified. As seen in Fig. 4A, BSPH1 is seen only in the pellet fraction (liposomes) following incubation and ultracentrifugation, indicating that the protein bound to the PC liposomes. As a positive control, we also tested the binding of crude BSP proteins (cBSP) to PC liposomes. The major protein of cBSP is BSP1 (previously named BSP-A1/-A2), which has a molecular weight of 15–16 kDa. As seen in Fig. 4A, there is a 15–16 kDa doublet in the pellet fraction, indicating that these proteins also bound to PC liposomes. On the other hand, BSA did not bind to PC-liposomes, as seen by its presence in the supernatant after incubation and ultracentrifugation. To rule out the possibility that binding of BSPH1 to PC liposomes could be mediated by thioredoxin, we expressed thioredoxin alone and incubated purified protein with PC-liposomes. As seen in Fig. 4A, thioredoxin alone does not bind to PC liposomes, since it is completely found in the supernatant after incubation with liposomes and ultracentrifugation.
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Binding to LDL
Another property of BSP proteins is that they bind to certain lipoproteins, notably LDL. It has been shown that in bovine, the interaction between BSP proteins and LDL from hen’s egg yolk present in extenders used for semen cryopreservation is essential for sperm protection against cold shock (Bergeron et al., 2004). We sought to determine whether recombinant BSPH1 also displayed this binding property. To test this, we incubated Trx-BSPH1 with LDL isolated from egg yolk, and then floated the lipoproteins by ultracentrifugation of the protein–lipoprotein solution in the presence of KBr added to achieve a density of 1.21 g/l. Using this technique, proteins that have bound to LDL will be found in the top portion of the tube following ultracentrifugation, whereas unbound proteins will be found in the middle and bottom fractions. As shown in Fig. 4B (top panel), we detected recombinant BSPH1 in the top layer after incubation with LDL and ultracentrifugation, signifying that some of the protein had indeed bound to LDL. A signal is also present in the lane corresponding to the bottom fraction, indicating that a portion of the protein did not bind to the lipoprotein. Knowing that the binding capacity of LDL for BSP proteins is very high (Manjunath et al., 2002), it is unlikely that the BSP protein binding sites on LDL were saturated. A portion of Trx-BSPH1 may not have bound to LDL due to incorrect folding or inaccessibility of the binding sites. As controls, Fig. 4B (middle and bottom panels) shows that Trx alone binds LDL very weakly, indicating that binding of Trx-BSPH1 to LDL is not due to the thioredoxin tag, and that the bovine BSP proteins bind LDL strongly.
Binding to human sperm
The most important binding property tested was binding to the sperm membrane. Recombinant BSPH1 was incubated with human sperm, after which sperm were washed and the sperm membrane proteins extracted. As seen in Fig. 4C (top panel), an immunoreactive band is observed in the lanes corresponding to extracted sperm proteins, while no reaction is seen in wash fractions. An intense band is observed in the lane corresponding to the supernatant after centrifugation of the sperm-recombinant protein mixture. This could signify that only a portion of our protein is correctly folded and retains sperm-binding activity, and/or that excess protein was added in comparison to the number of binding sites present on the quantity of sperm used for the experiment. Also, knowing that BSPH1 mRNA is expressed in the epididymis (Lefebvre et al., 2007), sperm are likely to come in contact with this protein during epididymal transit. Upon ejaculation, sperm are mixed with seminal plasma, a fluid containing numerous sperm-binding proteins that could mask BSPH1 binding sites. As a control, we incubated thioredoxin with human sperm and repeated the same procedure. As seen in Fig. 4C (middle panel), only a very faint signal is observed in the sperm-extracted proteins after incubation of sperm with thioredoxin, indicating that thioredoxin alone does not or very weakly binds to human sperm. In addition, untreated sperm incubated in the absence of recombinant protein (negative control) were analysed in the same way, and no immunoreaction was observed (Fig. 4C, bottom panel). As a positive control, the binding of cBSP proteins to human sperm was also verified. As expected, these proteins indeed bind to human sperm (data not shown). Thus, the BSPH1 protein produced as a thioredoxin fusion was soluble and active, as shown by its ability to bind PC liposomes, LDL and human sperm.
Despite several studies comparing the efficiency of different fusion tags (GST, maltose-binding protein, thioredoxin etc), the mechanism behind the solubilizing effect of fusion tags on target proteins remains unclear (Dyson et al., 2004; Hammarstrom et al., 2006). In the case of thioredoxin, it has been proposed that it could exert its effect either by acting as a covalently linked chaperone, aiding in protein folding, or as an oxidoreductase, which would catalyse the formation of correct disulfide bonds. A literature survey on thioredoxin-fused recombinant proteins suggests that proteins devoid of disulfide bonds can be successfully produced in the soluble form in E. coli cells bearing a reducing cytoplasm (ex. BL21) (Liu et al., 2007; Sun et al., 2007), suggesting a role for thioredoxin in protein folding. On the other hand, most proteins containing disulfide bridges require not only fusion to thioredoxin, but also expression in genetically modified E. coli strains having an oxidizing cytoplasm (ex. Origami B (DE3), BL21trxB (DE3), AD494 (DE3)), for soluble expression (Moura-da-Silva et al., 1999; Lauber et al., 2001; Lehmann et al., 2003; Xiong et al., 2005; Peisley and Gooley, 2007). This would suggest that thioredoxin could also be catalysing proper disulfide bond formation. However, in certain cases, proteins harbouring single or multiple disulfide bridges were successfully expressed in the soluble form in BL21 (DE3) cells (Zhao et al., 2000; Yuan and Hua, 2002; Cui et al., 2007). In the present study, His-tagged BSPH1 was not expressed in the soluble form in BL21 (DE3)pLysS cells, and only a slight improvement was seen with expression in Origami B (DE3)pLysS cells. However, fusion to thioredoxin dramatically improved soluble expression in Origami B (DE3)pLysS cells, with minimal improvement in BL21 (DE3)pLysS cells. This suggests that thioredoxin promotes the solubility of BSPH1 by catalysing proper disulfide bond formation, although it could also be acting as a molecular chaperone.
In conclusion, after a considerable amount of optimization and troubleshooting, an efficient method for recombinant expression of soluble BSPH1 has been developed. Recombinant BSPH1 protein was shown to be active by its ability to bind PC liposomes, LDL and human sperm, which reflects its potential biological function in sperm maturation, similarly to the bovine BSP proteins. The strategy developed here was also successful in producing recombinant murine Bsph1, strongly suggesting its application for all BSP-homologous proteins. In addition, this method could be used to express Fn2 domains contained in non BSP-related proteins. Purified recombinant BSPH1 will aid in establishing the biological role of this protein, a task that was previously impossible due to the unavailability of native protein. For example, purified recombinant BSPH1 will be added to washed ejaculated human sperm to study its effect on sperm capacitation, motility and viability. In addition, the possible role of this epididymal protein in promoting sperm-oocyte interaction will be investigated. Since BSP proteins are known to promote cholesterol and phospholipid efflux from bovine sperm during capacitation, the effect of recombinant proteins on lipid efflux will also be verified. In the same way, studies can be conducted using recombinant mouse Bsph1 and murine epididymal sperm. Since BSPH1 and Bsph1 are orthologues, results generated in mouse can be extrapolated to human. Studying the role of BSPH1 in human sperm functions may identify a new factor involved in human fertility, which can be beneficial for infertility diagnosis and for the future development of male contraceptives.
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TopAbstractIntroductionMaterials and MethodsResults and discussionFundingAcknowledgementsReferences |
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This work was funded by the Canadian Institutes of Health Research [MOP-86 510 to P.M.]. J.L. is a recipient of a Doctoral Training Award from the Fonds de la Recherche en Santé du Québec.
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Acknowledgements |
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TopAbstractIntroductionMaterials and MethodsResults and discussionFundingAcknowledgementsReferences |
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We thank Dr Yves Paquette for his advice in cloning and troubleshooting.
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Submitted on August 8, 2008; resubmitted on November 25, 2008; accepted on December 3, 2008.
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