PEDS Advance Access originally published online on December 10, 2007
Protein Engineering Design and Selection 2007 20(12):625-633; doi:10.1093/protein/gzm066
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Expression and enzymatic characterization of the soluble recombinant plasmepsin I from Plasmodium falciparum
1Department of Food Science, University of Guelph, Guelph, ON, Canada N1G 2W1 2Department of Food and Bioproduct Sciences, University of Saskatchewan, Saskatoon, SK, Canada S7N 5A8 3Department of Applied Biological Science, Faculty of Agriculture, Kagawa University, Kagawa, Japan
4 To whom correspondence should be addressed.E-mail: ryada{at}uoguelph.ca
 |
Abstract |
|---|
 Top Abstract IntroductionMethodsResults and discussionFundingAcknowledgementsReferences |
|---|
The plasmepsins are involved in the degradation of host cell hemoglobin during malaria infection. Plasmepsin I (PM I) initiates the degradative process, and has been suggested as an attractive target for the treatment of malaria. The production of active recombinant PM I, however, has been challenging. We report for the first time, the expression and partial characterization of soluble recombinant PM I from Plasmodium falciparum in which a truncated form of PM I (Lys77P–Leu329) (P indicates a propart residues) was fused to thioredoxin in the pET32b(+) vector, Trx-tPM I and expressed in Escherichia coli Rosetta-gami B (DE3)pLysS. The soluble fusion protein was purified from cell culture using a combination of Ni2+ affinity and gel filtration chromatography and was capable of autocatalytic activation at pH 4.0–5.5, which occurred at Leu116P–Ser117P, seven residues upstream of the native cleavage site (Gly123P–Asn1). The mature tPM I (mtPM I) was capable of hydrolyzing both human hemoglobin with a pH optimum of pH 2.8–4.0 and the synthetic fluorogenic peptide EDANS-CO-CH2-CH2-CO-ALERMFLSFP-Dap(DABCYL)-OH with a dual pH optima of pH 2.5–3.0 and pH 4.5–5.5. Using the synthetic substrate, mtPM I exhibited kinetic parameters comparable to native PM I.
Keywords: aspartic protease/malaria/plasmepsin I/recombinant expression
|
Introduction |
|---|
|
TopAbstractIntroductionMethodsResults and discussionFundingAcknowledgementsReferences |
|---|
Malaria afflicts up to several hundred million people resulting in the death of approximately 1–2 million (Greenwood et al., 2005
). The disease is caused by parasites of the genus Plasmodium. During P. falciparum infection, the most lethal of the four parasites infecting humans, the parasite invades erythrocytes and degrades large amounts of host cell hemoglobin in food vacuole via aspartic, cysteine and metallopeptidases (Kolakovich et al., 1997). The development of P. falciparum resistance to drugs such as chloroquine and sulphadoxine/pyrimethamine is a major challenge for malaria control; therefore, new treatment methods and alternative drugs are needed (Ridley, 2002). The plasmepsins produced by the Plasmodium parasite are aspartic proteases and have been recognized as an attractive target for the design of novel chemotherapeutic compounds for the control of malaria (Olliaro and Goldberg, 1995; Silva et al., 1996; Asojo et al., 2002; Boss et al., 2003; Johansson et al., 2004; Ersmark et al., 2006).
There are 10 plasmepsins, type II membrane proteins, reported in the genome of P. falciparum, PM I, II and IV–X and histo-aspartic protease (HAP) (Coombs et al., 2001). Four of them, PM I, PM II, HAP and PM IV, lie in a cluster on chromosome 14, and share 50–79% amino acid sequence identity (Berry et al., 1999). These four plasmepsins have been localized in the food vacuole of the parasite, and have been shown to be involved in hemoglobin degradation (Coombs et al., 2001; Banerjee et al., 2002). PM I and II appear to initiate the degradative process by cleaving a conserved hinge region in the 
-chain of native hemoglobin (Gluzman et al., 1994). HAP, PM IV and falcipain-2 and -3 subsequently act on the denatured or fragmented globin (Salas et al., 1995; Francis et al., 1996; Berry et al., 1999; Humphreys et al., 1999; Sijwali et al., 2001; Banerjee et al., 2002). PM V was shown to be localized in the endoplasmic reticulum and was suggested to have a role in intraerythrocytic biology that is distinct from PM I, PM II, HAP and PM IV (Klemba and Goldberg, 2005). PM I, II and IV are known to be classical aspartic proteases containing two catalytic aspartic acids, whereas one of the catalytic aspartic acids is replaced with a histidine in HAP (Berry et al., 1999; Coombs et al., 2001). To date, much of our knowledge of recombinant expression, crystallographic analysis, kinetics and inhibitor development comes mainly from the successful expression of PM II and IV in Escherichia coli (Hill et al., 1994; Silva et al., 1996; Asojo et al., 2002; Li et al., 2004; Clemente et al., 2006; Gutierrez-de-Teran et al., 2006; Kim et al., 2006). Until recently, the recombinant expression of HAP has been unsuccessful due to insolubility of the enzyme (Berry et al., 1999; Banerjee et al., 2002); however, our group was the first to successfully express, and partially characterize, soluble recombinant HAP as a thioredoxin fusion protein from E. coli (Xiao et al., 2006). Expression of active recombinant PM I has also been a major challenge (Coombs et al., 2001; Ersmark et al., 2006). Luker et al. (1996) found that recombinant PM I did not resemble its native homolog kinetically and was unable to degrade hemoglobin due to its inability to autoactivate. Moon et al. (1997) obtained active plasmepsin I by introducing a Lys110P
Val (P indicates a propart residues) mutation into the propart of the zymogen, but found that the overall efficiency of refolding and autoactivation remained low. In a further study, Tyas et al. (1999) reported a 3–10-fold diminution in the kcat values for substrate hydrolysis by this mutant, as compared with the native PM I. Although attempts to crystallize recombinant PM I have been made (Moon et al., 1997), no crystal structure has been reported to date. Furthermore, a large proportion of the inhibitors reported recently have not been assessed regarding PM I inhibition due to the difficulties in obtaining active enzyme (Ersmark et al., 2006). In the present work, we report the successful expression and partial characterization of active recombinant PM I using the strategy to express HAP (Xiao et al., 2006).
|
Methods |
|---|
|
TopAbstractIntroductionMethodsResults and discussionFundingAcknowledgementsReferences |
|---|
Cloning of PM I and construction of expression plasmid
The method for cloning of plasmepsin I (PM I) was similar to that used for HAP (Xiao et al., 2006). The PM I gene was isolated from the genomic DNA of P. falciparum strain 3D7 (MR4/American Type Culture Collection, Manassas, VA, USA) using polymerase chain reaction (PCR) with primers PMIFW (5'-AAGGATCCCATGGCTTTATCAATTAAAG-3') and PMIRV (5'-AATGGATCCTTACAATTTTTTTGGC-3'). The PCR products were cloned into vector pUC-19 to produce the plasmid construct pUC19–PM I. The sequence of the PM I gene was confirmed by sequencing with M13 primers using Dye Terminator Cycle Sequencing on an ABI PRISM Model 377 at the Guelph Molecular SuperCenter, University of Guelph. The transmembrane region of the propeptide was then deleted by amplification from pUC19-PM I using primer PMI-H227 (5'-CGGGATCCAAAACATGTAATAATTGG-3') and PMIRV to yield the truncated PM I (tPM I) (Lys77P–Leu329). The amplified fragment (tPM I) was digested with BamHI, and cloned into pET-32b(+) to give rise to pET32b-tPM I in which tPM I was fused to thioredoxin. The resulting construct plasmid encoded thioredoxin, followed by Tags including thrombin cleavage site, His-Tag, S-Tag and enterokinase cleavage site, and the truncated PM I (Lys77P–Leu329) (Fig. 1). The DNA sequence of the expression construct was confirmed by sequencing with T7 promoter and T7 terminator primers.
|
Expression of the fusion protein
Escherichia coli Rosetta-gami B (DE3)pLysS cells were transformed with the expression construct pET32b–tPM I. Transformants were cultured in 1 l of LB medium containing 12.5 µg/ml tetracycline, 15 µg/ml kanamycin, 34 µg/ml chloramphenicol and 50 µg/ml ampicillin until the OD600 reached 1.0 at 37°C, and then induced with 1 mM isopropyl-ß-D-thiogalactopyranoside (IPTG) for 3–4 h at 30°C as suggested by Xiao et al. (2006). Cells were collected by centrifugation (2500xg for 15 min).
Purification of the fusion protein
Cell pellets were resuspended in BugBusterTM reagent (Novagen, Madison, WI, USA) and incubated at room temperature for 20 min with gentle shaking. The sample was then centrifuged at 16 000xg for 20 min at 4°C. The supernatant and the insoluble cell debris were subsequently collected and analyzed using SDS–PAGE and western blot with anti-thioredoxin primary antibody (Invitrogen, Burlington, ON, Canada). The soluble fraction obtained above was applied on a HisSelectTM Cartridge (6.4 ml) (Sigma-Aldrich, Oakville, ON, Canada); the column was first washed with 50 mM sodium phosphate/0.3 M NaCl/10 mM imidazole pH 7.5 washing buffer and then with 10% elution buffer (50 mM sodium phosphate/0.3 M NaCl/250 mM imidazole pH 7.5) and finally eluted with 100% elution buffer. The sample was further purified by gel filtration with a SuperoseTM 12 10/300 GL (GE Healthcare, Uppsala, Sweden) column equilibrated with 50 mM sodium phosphate/150 mM NaCl pH 7.5 to obtain the fusion Trx-tPM I protein. Purification was monitored using SDS–PAGE and western blot with anti-PM I antibody (MR4/American Type Culture Collection, Manassas, VA, USA).
Activation, purification and N-terminal sequenceof the active recombinant PM I
In order to determine the pH optimum for activation of PM I, 2 µg of purified thioredoxin-tPM I (Trx-tPM I) was incubated in 100 mM sodium acetate at various pH values (pH 3.5–6.5) at 37°C for 4 h. For the time course activation, 2 µg of Trx-tPM I was incubated in 100 mM sodium acetate, pH 4.5, for 15 min, 30 min, 1 h, 2 h and 4 h. To examine the effect of protease inhibitors on the autocatalytic processing of PM I, 2 µg of Trx-tPM I was incubated with or without the aspartic protease inhibitor pepstatin A (10 µM), cysteine protease inhibitor N-acetyl-Leu-Leu-norleucinal (ALLN, 15 µM), and serine protease inhibitors PMSF (1 mM) and leupeptin (100 µM) in 100 mM sodium acetate, pH 4.5, at 37°C for 4 h (Kim et al., 2006). The processing of PM I was detected by SDS–PAGE.
The activated sample at pH 4.5 was further purified by washing in an Ultracel YM50 Centricon (Millipore, Billerica, MA, USA) to remove the cleaved prosegment and thioredoxin to obtain mature tPM I (mtPM I). mtPM I protein was subjected to SDS–PAGE and electroblotted to a polyvinylidene difluoride (PVDF) membrane for N-terminal sequence analysis by the Advanced Protein Technology Center at the Hospital for Sick Children (Toronto, ON, Canada).
Protein concentration determinations
Enzyme and protein concentrations were determined in triplicate by the DC protein Assay (Bio-Rad, Hercules, CA, USA) using bovine serum albumin as the standard (Lowry et al., 1951).
SDS–PAGE was performed according to the method of Laemmli (1970) in a Mini-Protean II electrophoresis cell (Bio-Rad, Hercules, CA, USA). Protein samples were mixed with sample buffer in the presence of β-mercaptoethanol and SDS, heated for 5 min at 100°C, and loaded onto a 15% polyacrylamide gel with SDS. Gels were run at a constant voltage (200 V) for 1 h. The separated proteins were either stained with Coomassie blue or transferred onto PVDF membrane (Bio-Rad, Hercules, CA, USA) for western blotting according to the method of Towbin et al. (1979) using either anti-thioredoxin antibody or anti-PM I antibody. Electroblotting was carried out for 1 h at a constant voltage (100 V, 4°C) on a Mini Transblot electrophoretict transfer cell (Bio-Rad, Hercules, CA, USA).
Native PAGE was carried out similar to SDS–PAGE except that the sample buffer contained neither SDS nor β-mercaptoethanol (samples were not heated), and the 10% polyacrymide gel and electrophoresis solutions were prepared without SDS. Gels were run at a constant voltage (80 V) for 2.5 h. The gels were developed with GelCode Blue Stain Reagent (Pierce, Rockford, IL, USA).
The hemoglobin zymography gel was prepared similar to native PAGE except 0.8 mg of human hemoglobin (dissolved in 100 mM acetic acid-HCl, pH 2.0) was co-polymerized in 5 ml of 10% polyacrylamide. After electrophoresis, the gels were incubated in 100 mM sodium acetate buffer (pH 2.8 or 5.0) at 37°C for 2.5 h, and then developed with GelCode Blue Stain Reagent.
The pH-optimum of mtPM I was determined using the internally quenched fluorescent synthetic peptide substrate EDANS-CO-CH2-CH2-CO-Ala-Leu-Glu-Arg-Met-Phe-Leu-Ser-Phe-Pro-Dap-(DABCYL)-OH (2837b) with a cleavage site of Phe-Leu (AnaSpec Inc., San Jose, CA, USA), previously described by Istvan and Goldberg (2005), at various pH values between 2.0 and 6.5 in 100 mM sodium acetate buffers (pH 2.0–3.0 acetic acid solutions were adjusted with hydrochloric acid). 3 nM mtPM I (37.8 kDa) and 1.5 µM substrate were used for the assays performed at 25°C. The assays were carried out using a Victor 2 1420 Multilabel Counter (Perkin Elmer, Woodbridge, ON, Canada) with excitation at 335 nm and emission at 500 nm (Istvan and Goldberg, 2005).
The sequence of mature PM I was submitted to the Geno3D server at the Institute for the Biology and Chemistry of Proteins, University of Lyon, France (Combet et al., 2002) and a predicted molecular model was created. This model was then energy-minimized using the NAMD molecular modeling program (Kale et al., 1999). The calculation used topology force field data provided with the package, and was carried out in a water-filled box. The parameters were adjusted to a 15-Å cut-off distance and for 10 000 runs.
In order to determine the hemoglobin degradation by PM I, 5 µg of human hemoglobin (Sigma-Aldrich, Oakville, ON, Canada) were incubated with 10 nM mtPM I in 100 mM sodium acetate at various pH values (pH 2.0–6.5, pH 2.0–3.0 acetic acid solutions were adjusted with hydrochloric acid) for 6 h. The effect of protease inhibitors on the hemoglobin degradation by PM I was assessed by incubating 5 µg of hemoglobin with 10 nM mtPM I in 100 mM sodium acetate (pH 2.8) in the presence or absence of pepstatin A (10 µM) or ALLN (15 µM) for 3 and 6 h at 37°C. The reaction was terminated by heating at 95°C in sample buffer containing 1.2% SDS. Hemoglobin degradation was detected by SDS–PAGE (Xiao et al., 2006).
Enzyme kinetic assays and inhibition studies
Kinetic parameters were determined using substrate 2837b as described above. The reaction was carried out at 25°C in 100 mM sodium acetate pH 2.8 and 5.0 using 3 nM mtPM I and 0.1 µM–4.0 µM substrate. The initial reaction rates were determined by calculating the slope of the linear portion of the curve (fluorescence/min). The observed fluorescence units were converted to micromoles per minute using a conversion factor derived from a standard curve for the complete digestion of the substrate by commercial yeast proteinase A (Sigma-Aldrich, Oakville, ON, Canada) at four substrate concentrations (0.1, 1.0, 2.0 and 5.0 µM) (Xiao et al., 2006). Non-linear regression of the Michaelis-Menten model was used to derive Km, and kcat which were calculated from kcat=Vmax/[E]. The concentration of active enzyme [E] used in each assay was determined by active site titration with pepstatin A. Inhibition assays of mtPM I were done with pepstatin A at various concentrations (0.16–40 nM) and ALLN (5–160 µM). The reaction was carried out in 100 mM sodium acetate pH 2.8 and 5.0 using 3 nM mtPM I and 1.5 µM peptide substrate. Ki values for pepstatin A and ALLN were determined with GraphPad Prism (Version 3.03) using the equation of Cheng and Prusoff (1973).
|
Results and discussion |
|---|
|
TopAbstractIntroductionMethodsResults and discussionFundingAcknowledgementsReferences |
|---|
Soluble expression of recombinant plasmepsin I in E. coli
Similar to the protocol used to express HAP (Xiao et al., 2006), a dual-approach was used for PM I where the truncated PM I (K77P–L329) combined with the deletion of the membrane-binding domain was fused with the highly soluble protein, thioredoxin (Trx-tHAP) in the pET-32b(+) vector and expressed in E. coli Rosetta-gami B (DE3)pLysS. As observed for recombinant HAP (Xiao et al., 2006), a small proportion of total tPM I was soluble in the cell lysate supernatant as illustrated in SDS–PAGE (Fig. 2A, lane 1) and the western blot with anti-thioredoxin antibody (Fig. 2B, lane 1). Eight percent of the total Trx-tPM I fusion protein was expressed in the soluble form as determined by densitometric analysis of a Coomassie stained SDS–PAGE gel using Syngene GeneToolsTM software (data not shown). In contrast, about 13% of the total fusion protein was expressed in a soluble form for Trx-tHAP (Xiao et al., 2006) indicating that PM I was more difficult to express in vitro than HAP.
|
Purification of Trx-tPM I from the cell lysate supernatant was accomplished by FPLC using a combination of Ni2+ affinity and size exclusion chromatography (Fig. 2C and D). The supernatant was initially applied on a HisSelectTM column on an AKTA FPLC system (GE Healthcare, Uppsala, Sweden). The elution was further purified by gel filtration with a SuperoseTM 12 10/300 GL column to obtain the fusion Trx-tPM I protein (Fig. 2C and D). The purification steps are shown from SDS–PAGE (Fig. 2C) and western blot using anti-PM I antibody (Fig. 2D) and summarized in Table I. This purification procedure produced 62 µg of Trx-tPM I fusion protein from 1 l of cell culture, which could be activated by acidification to the active mature form (Fig. 3).
|
|
Autocatalytic processing of PM I
The recombinant Trx-tPM I fusion protein was incubated in 100 mM buffer over the pH range 3.5–6.5 at 37°C for 4 h. The fusion protein (Trx-tPM I) underwent activation between pH 4.0 and 5.5 with a pH optimum of 4.5 (Fig. 3A, lane 4), as evidenced by the band shifting from 60.3 kDa (F) to 37.8 kDa (M), which is similar to the reported size of mature form of PM I (Banerjee et al., 2003). The pH optimum of activation was similar to that reported for non-recombinant native PM I, pH 4.0–5.5 (Banerjee et al., 2003). The time course study showed that hydrolysis for 2 h at 37°C at pH4.5 was sufficient to convert fusion protein to the mature form (Fig. 3B). There were intermediate products, intermediate Trx-tPM I (identified as I) and tPM I (identified as Z) produced during the activation (Fig. 3). N-terminal sequencing showed that a cleavage between Met and Asp in the region of the Tags (Fig. 1) yielded an intermediate Trx-tPM I (I) with a molecular mass of 44.5 kDa, and a cleavage between Phe83P and Ser84P which resulted in tPM I (Z) with a molecular mass of 41.8 kDa (Figs 1 and 3). It would appear that the activation process proceeded in three steps: from fusion protein (Trx-PM I) to intermediate Trx-tPM I (I), from intermediate Trx-tPM I (I) to truncated zymogen tPM I (Z) and finally from tPM I (Z) to mature PM I (M) (Figs 1 and 3).
After incubation at pH 4.5 for 4 h, the resultant protein was washed in an Ultracel YM50 Centricon to remove thioredoxin and the cleaved prosegment to obtain the mature enzyme (mtPM I) (Fig. 4A). N-terminal sequencing of the purified mtPM I (Fig. 4A) confirmed the identity of mtPM I and showed that autocatalytic cleavage occurred between amino acids Leu116P and Thr117P, seven residues upstream of the native cleavage site at Gly123P–Asn1 identified by Gluzman et al. (1994) (Fig. 1). Moon et al. (1997), in a study which mutated Lys110P to Val110P at the propart of the zymogen, reported that there were two autocatalytic cleavage sites for an activation site mutant of the PM I; the first one was 12 residues upstream of the native cleavage site and the second one is the same as reported in the present study. Tyas et al. (1999), who also mutated Lys110PVal, reported only the first cleavage site.
|
In order to determine which protease inhibitors could prevent the autocatalytic processing of PM I, the Trx-tPM I fusion protein was incubated with various protease inhibitors in 100 mM sodium acetate (pH 4.5) for 4 h at 37°C. The results showed that the processing of PM I was completely inhibited by aspartic protease inihibitor pepstatin A (Fig. 3C, lane 3). The cysteine protease inhibitor ALLN showed some inhibition (Fig. 3C, lane 4), while the serine protease inhibitor PMSF and leupeptin were not inhibitory (Fig. 3C, lanes 5 and 6). It has been reported that in vivo, pepstatin A does not inhibit proplasmepsin processing, whereas ALLN completely inhibits proplasmepsin processing (Francis et al., 1997
; Banerjee et al., 2003). The fact that ALLN did not inhibit the processing of PM I (Fig. 3C, lane 4), PM II (Kim et al., 2006) and PM VI (Kim et al., 2006) in vitro, but did so in vivo (Francis et al., 1997; Banerjee et al., 2003) and that the activation sites of PM I, PM II and PM VI in vitro were different from those in vivo (Fig. 1; Tyas et al., 1999; Banerjee et al., 2003; Li et al., 2004) may support the suggestion that intracellular, proplasmepsin processing is not autocatalytic but is mediated by an atypical calpain-like protease (Francis et al., 1997; Banerjee et al., 2003). Kim et al. (2006), however, suggested that the processing of both PM II and IV in vivo occurred by autocatalytic and transcatalytic events between them, rather than by other proteases. pH dependence of mature enzyme activity
The pH-activity profile for mtPM I was determined over the pH range 2.0–6.5 using internally quenched fluorescent peptide substrate EDANS-CO-CH2-CH2-CO-Ala-Leu-Glu-Arg-Met-Phe-Leu-Ser-Phe-Pro-Dap-(DABCYL)-OH (2837b) (Fig. 4B). One hour activated mtPM I was shown to have two pH optima at pH 4.5–5.5 and pH 2.5–3.0. The first optimum (pH 4.5–5.5) is similar to that previously reported for the native PM I, i.e. pH 4.5–5.0 (Goldberg et al., 1991) and to that of the food vacuole (
pH 5.0) (Yayon et al., 1984); however, it became less prominent when the time of autoactivation at 37°C was greater than 4 h (Fig. 4B). Hydrolysis of human hemoglobin with 4 h activated mtPM I confirmed that mtPM I effectively degraded the native hemoglobin in 6 h and the degradation was faster at low pH range (pH 2.8–4.0) (Fig. 4C). Interestingly, the 1 h-activated mtPM I also had maximal activity at pH 2.8–4.0 in the degradation of hemoglobin (Fig. 4D), although it showed high activity at pH 4.5–5.5 on the synthetic peptide substrate (Fig. 4B). This indicated that the recombinant mtPM I had a different pH optima of activity on the peptide and protein substrate.
The pH dependency of mtPM I was further examined with native-, SDS–PAGE and zymography. It was shown that mtPM I consisted of two forms when it was analyzed using native PAGE (Fig. 5A, lane 1), which was supported by size exclusion chromatography data where there was both a broad peak near the void volume and a peak eluting later suggesting that mtPM I contained both oligomeric (aggregates) and monomeric forms, respectively (data not shown). The treatment of mtPM I with 0.2% CHAPS, zwitterionic surfactant, reduced the amount of aggregates with a corresponding increase in the amount of monomers as shown in Fig. 5A, lane 2, which coincided with an increase in activity at pH 5.0 and a decrease at pH 2.8 (Fig. 5B). These observations suggested that the monomer is more active at pH 5.0 while the aggregates are more active at pH 2.8. This suggestion was confirmed through further experiments. The activation of PM I at 4°C (A1) and 37°C (A2) gave similar amounts of the mature form shown in the SDS–PAGE gel (Fig. 6A). However, A1 sample contained a higher amount of monomers in contrast to A2 sample as indicated in the Native PAGE (Fig. 6B). Correspondingly, the activity of A1 sample at pH 5.0 was more pronounced than that of A2 sample at pH 5.0 (Fig. 6E). Native PAGE also indicated that the higher activity of A1 at pH 2.8 as compared to A2 at pH 2.8 paralleled the higher amount of aggregates in A1 than that in A2 (Fig. 6B and E). In addition, A1, which had more monomers, showed a higher activity at pH 5.0 than at pH 2.8, while A2, which consisted of more aggregates, showed a higher activity at pH 2.8 than at pH 5.0 (Fig. 6B and E). The activity of the aggregate form of mtPM I was further illustrated using hemoglobin zymography, in which the aggregates in both A1 and A2 could degrade hemoglobin and the degradation was faster at pH 2.8 than at pH 5.0 (Fig. 6C). These results clearly indicated that the monomers and aggregates of PM I had a different pH optimum, with monomers at pH 4.5–5.5 and the aggregates at pH 2.5–3.0, and thus the observed dual pH optima described above resulted from the existence of two states of the mature enzymes (i.e. monomeric form and aggregate form). Different oligomeric states with different pH optimum have been reported for various enzymes (Campillo et al., 1981; Wong et al., 1987; Klyachko et al., 2005). For example, Campillo et al. (1981) found that the tetrameric - galactosidase have a pH optimum of about pH 7.0, whereas the monomeric form display a pH optimum of 5.6.
|
|
Further analysis also revealed that the monomers and aggregates of PM I had different substrate specificities on synthetic peptide and human hemoglobin. As shown in Fig. 6D, both monomer-dominant A1 and aggregate-dominant A2 samples showed higher activities to human hemoglobin and BSA at pH 2.8 than at pH 5.0. If the monomeric form hydrolyzes hemoglobin efficiently, then the monomer-dominant A1 should degrade hemoglobin faster at pH 5.0 than at pH 2.8, since it exhibited higher activities towards the synthetic peptide 2837b at pH 5.0 compared to pH 2.8 (Fig. 6E, A1). However, the opposite was observed, i.e. the monomer-dominant A1 degraded hemoglobin faster at pH 2.8 than at pH 5.0. Therefore, it was suggested that the monomers and aggregates of PM I hydrolyze synthetic peptide and human hemoglobin differently. The aggregates were able to hydrolyze both proteins and peptides with similar efficiency while the monomers were more efficient at hydrolyzing the synthetic peptides as compared to proteins. It was reported that all plasmepsin structures determined, so far, are tightly associated homodimers (Asojo et al., 2003
). The observation that aggregates and monomers hydrolyze proteins and peptides with different efficiencies may have biological relevance in vivo.
The dual pH optima and the possible mechanism of soluble aggregates for mtPM I in hemoglobin degradation were further investigated by molecular modeling. The molecular modeling of PM I gave similar structural models to PM II and other aspartic proteases (Fig. 7) (Asojo et al., 2003). The examination of the structure models of PM-I and other aspartic protease models revealed that the active site clefts have little difference in the residues that can be affected by pH changes around pH 4.0, i.e. Asp or Glu. There is an Asp residue at position 303 near the active centre, Asp34 and Asp214. However, Asp303 was conserved among the most aspartic proteases that exhibit single pH optima (Fruton, 1971). In other words, Asp303 is not likely to be the cause of the double pH optima observed in this PM I. According to Asojo et al. (2003), PM II has a unique and close relation between two monomers in the crystals that showed an unusual P2 space group. They suggested that PM II has the ability to form a dimeric structure in the solution unlike other aspartic proteases, which suggests that PM I can form a dimeric structure similar to PM II since a BLAST search showed that PM I shares 80% amino acid sequence homology with other plasmepsins (PM II, PM IV, HAP) while only 52% or less with the most of other aspartic proteases (such as rennin, cathepsin D). The model of PM I was, therefore, examined to see if dimer formation can be affected by pH of the environment. The assumed interface of two monomers had two pairs of salt bridges in PM I (Fig. 8) whereas PM II (PDB: 1IL3) had one pair. Asp107–Lys238 existed in both enzymes and this Lys238 is on the flexible loop that protrudes into the active site cleft of the other monomer. This flexible loop would have the similar local structures between PM I and PM II. The other pair, Lys53–Asp279 was observed only in PM I. Between pH 2.0 and 4.0, which is below the pKa of free Asp (pH 3.9), Asp279 is likely protonated and loses the ability to form the salt bridge. As a result, the mode of interaction changes from what exists in the monomer–monomer interaction in the pH 4.0–6.0 range leaving Lys53 charged without pairing to a neutralizing residue, i.e. inability to form a salt bridge with another Asp or Glu which are abundant on the surface of PM I. We speculate that this situation will initiate the aggregation of PM I in the pH 2.0–4.0 range. The formation of salt bridges among aggregated monomers is consistent with the observation in the present study where CHAPS treatment reduced the amount of aggregated monomers (Fig. 5A). This formation of aggregates would involve the change of environments of active site clefts in the aggregated PM I and result in the pH optima shift from pH 5.0 to pH 2.8.
|
|
The degradation of hemoglobin by mtPM I could be inhibited by pepstatin A, but not by ALLN (Fig. 9). Kim et al. (2006)
reported that hemoglobin degradation by PM II or IV was completely inhibited by 10 µM pepstatin A, and PM II-mediated hemoglobin degradation was partially inhibited by 15 µM ALLN, while the same concentration of ALLN completely blocked hemoglobin degradation by PM IV. The above results indicate that homologous plasmepsins have different affinities for ALLN.
|
Kinetic analysis of mature PM I
Kinetic parameters of mtPM I for cleavage of the Phe33–Leu34 peptide bond in -globin at the optimum pH of 2.8 and 5.0 were determined using the quenched fluorescent peptide substrate 2837b previously used for the kinetic characterization of PM II (Istvan and Goldberg, 2005) and HAP (Xiao et al., 2006). The amount of active mtPM I was titrated with pepstatin A, and the kinetic data (Table II) were compared with data reported for native PM I from Luker et al. (1996), who also used the similar quenched fluorescent peptide substrate. mtPM I had a 1.3-fold higher Km and 0.3-fold lower kcat at pH 5.0 than at pH 2.8, which was 69% less effective in the hydrolysis of the peptide substrate at pH 5.0 than at pH 2.8. mtPM I at pH 2.8 had a Km of 0.87 µM and a kcat of 1.7 s–1 that were comparable to the kinetic parameters observed for native PM I at pH 5.0 (Table II) (Luker et al., 1996). mtPM I bound pepstatin A was similar to native PM I. However, as compared to pepstatin A, mtPM I had a very low affinity for ALLN, which may explain why the degradation of native hemoglobin was not inhibited by ALLN (15 µM) (Fig. 9).
|
In conclusion, the above results showed that the properties of recombinant PM I were generally comparable to native PM I (Table II, Luker et al., 1996
). The finding that the aggregates of the recombinant PM I was active is significant and deserves further investigation. The ability to utilize an expression system that combines a simple activation protocol with a relatively easy purification and produces sufficient amounts of protein, will undoubtedly allow for comprehensive structure-function studies of PM I to be undertaken which may lead to the potential development of specific inhibitors. |
Funding |
|---|
|
TopAbstractIntroductionMethodsResults and discussionFundingAcknowledgementsReferences |
|---|
This research is supported by the Natural Sciences and Engineering Research Council of Canada and the Canada Research Chair Program.
|
Footnotes |
|---|
Edited by Mirek Cygler
|
Acknowledgements |
|---|
|
TopAbstractIntroductionMethodsResults and discussionFundingAcknowledgementsReferences |
|---|
We are grateful to Derek Dee for his insightful comments.
|
References |
|---|
|
TopAbstractIntroductionMethodsResults and discussionFundingAcknowledgementsReferences |
|---|
Asojo O.A., Afonina E., Gulnik S.V., Yu B., Erickson J.W., Randad R., Medjahed D., Silva A.M. Acta Cryst. (2002) 58:2001–2008.[Web of Science]
Asojo O.A., Gulnik S.V., Afonina E., Yu B., Ellman J.A., Haque T.S., Silva A.M. J. Mol. Biol. (2003) 327:173–181.[CrossRef][Web of Science][Medline]
Banerjee R., Liu J., Beaty W., Pelosof L., Klemba M., Goldberg D.E. Proc. Natl Acad. Sci. USA (2002) 99:990–995.
Banerjee R., Francis S.E., Goldberg D.E. Mol. Biochem. Parasitol. (2003) 129:157–165.[CrossRef][Web of Science][Medline]
Berry C., Humphreys M.J., Matharu P., Granger R., Horrocks P., Moon R.P., Certa U., Ridley R.G., Bur D., Kay J. FEBS Lett. (1999) 447:149–154.[CrossRef][Web of Science][Medline]
Boss C., Richard-Bildstein S., Weller T., Fischli W., Meyer S., Binkert C. Curr. Med. Chem. (2003) 10:883–907.[CrossRef][Web of Science][Medline]
Campillo E.D., Shannon L.M., Hankins C.N. J. Biol. Chem. (1981) 256:7177–7180.
Cheng Y., Prusoff W.H. Biochem. Pharmacol. (1973) 22:3099–3108.[CrossRef][Web of Science][Medline]
Clemente J.C., et al. Acta Crystallogr. D. Biol. Crystallogr. (2006) 62:246–252.[CrossRef][Medline]
Combet C., Jambon M., Deléage G., Geourjon C. Bioinformatics (2002) 18:213–214.
Coombs G.H., Goldberg D.E., Klemba M., Berry C., Kay J., Mottram J.C. Trends Parasitol. (2001) 17:532–537.[CrossRef][Web of Science][Medline]
Ersmark K., Samuelsson B., Hallberg A. Med. Res. Rev. (2006) 26:626–666.[CrossRef][Web of Science][Medline]
Francis S.E., Gluzman I.Y., Oksman A., Banerjee D., Goldberg D.E. Mol. Biochem. Parasitol. (1996) 83:189–200.[CrossRef][Web of Science][Medline]
Francis S.E., Banerjee R., Goldberg D.E. J. Biol. Chem. (1997) 272:14961–14968.
Fruton J.S. The Enzyme—Boyer P.D., ed. (1971) New York: Academic Press. 119–164.
Gluzman I.Y., Francis S.E., Oksman A., Smith C.E., Duffin K.L., Goldberg D.E. J. Clin. Invest. (1994) 93:1602–1608.[Web of Science][Medline]
Goldberg D.E., Slater A.F., Beavis R., Chait B., Cerami A., Herderson G.B. J. Exp. Med. (1991) 173:961–969.
Greenwood B.M., Bojang K., Whitty C.J., Targett G.A. Lancet (2005) 365:1487–1498.[CrossRef][Web of Science][Medline]
Gutierrez-de-Teran H., Nervall M., Ersmark K., Liu P., Janka L.K., Bunn B., Hallberg A., Aqvis J. Biochemistry (2006) 45:10529–10541.[CrossRef][Web of Science][Medline]
Hill J., Tyas L., Phylip L.H., Dunn B.M., Berry C. FEBS Lett. (1994) 26:155–158.
Humphreys M.J., Moon R.P., Klinder A., Fowler S.D., Rupp K., Bur D., Ridley R.G., Berry C. FEBS Lett. (1999) 463:43–48.[CrossRef][Web of Science][Medline]
Istvan E.S., Goldberg D.E. J. Biol. Chem. (2005) 280:6890–6896.
Johansson P.O., et al. J. Med. Chem. (2004) 47:3353–3366.[CrossRef][Web of Science][Medline]
Kale L., Skeel R., Bhandarkar M., Brunner R., Gursoy A., Krawetz N., Phillips J., Shinozaki A., Varadarajan K., Schulten K. J. Comp. Phys. (1999) 151:283–312.[CrossRef]
Kim Y.M., Lee M.H., Piao T.G., Lee J.W., Kim J.H., Lee S., Choi K.M., Jiang J.H., Kim T.U., Park H. J. Biochem. (2006) 139:189–195.
Klemba M., Goldberg D.E. Mol. Biochem. Parasitol. (2005) 143:183–191.[CrossRef][Web of Science][Medline]
Klyachko N.L., Shchedrina V.A., Efimov A.V., Kazakov S.V., Gazayan I.G., Kristal B.S., Brown A.M. J. Biol. Chem. (2005) 280:16106–16114.
Kolakovich K.A., Gluzman I.Y., Duffin K.L., Goldberg D.E. Mol. Biochem. Parasitol. (1997) 87:123–135.[CrossRef][Web of Science][Medline]
Laemmli U.K. Nature (1970) 227:680–685.[CrossRef][Medline]
Li T., Yowell C.A., Beyer B.B., Hung S.-H., Westling J., Lam M.T., Dunn B.M., Dame J.B. Mol. Biochem. Parasitol. (2004) 135:101–109.[CrossRef][Web of Science][Medline]
Lowry O.H., Rosebrough N.J., Farr A.L., Randall R.J. J. Biol. Chem. (1951) 193:265–275.
Luker K.E., Francis S.E., Gluzman I.Y., Goldberg D.E. Mol. Biochem. Parasitol. (1996) 79:71–78.[CrossRef][Web of Science][Medline]
Moon R.P., et al. Eur. J. Biochem. (1997) 244:552–560.[Web of Science][Medline]
Olliaro P.L., Goldberg D.E. Parasitol. Today (1995) 11:294–297.[CrossRef][Web of Science][Medline]
Ridley R.G. Nature (2002) 415:686–693.[CrossRef][Medline]
Salas F., Fichmann J., Lee G.K., Scott M.D., Rosenthal P.J. Infect. Immun. (1995) 63:2120–2125.[Abstract]
Sijwali P.S., Shenai B.R., Gut J., Singh A., Rosenthal P.J. Biochem. J. (2001) 360:481–489.[CrossRef][Web of Science][Medline]
Silva A.M., et al. Proc. Natl Acad. Sci. USA (1996) 93:10034–10039.
Towbin H., Staehelin T., Gordon J. Proc. Natl Acad. Sci. USA (1979) 76:4350–4354.
Tyas L., Gluzman I., Moon R.P., Rupp K., Westling J., Ridley R.G., Kay J., Goldberg D.E., Berry C. FEBS Lett. (1999) 454:210–214.[CrossRef][Web of Science][Medline]
Wong J.H., Yee B.C., Buchanan B.B. J. Biol. Chem. (1987) 262:3185–3191.
Xiao H., Sinkovits A.F., Bryksa B.C., Ogawa M., Yada R.Y. Protein Expr. Purif. (2006) 49:88–94.[CrossRef][Web of Science][Medline]
Yayon A., Cabantchik Z.I., Ginsburg H. EMBO J. (1984) 3:2695–2700.[Web of Science][Medline]
Received January 28, 2007; revised October 16, 2007; accepted October 23, 2007.
CiteULike 
Connotea 
Del.icio.us What's this?
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||