Journal of Undergraduate Research
Volume 6, Issue # - Month Year

Refolding and Purification of “short” recombinant human Cathepsin D and Variants

Ambar Patel

ABSTRACT

Human cathepsin D (hCatD) is an aspartic peptidase synthesized at neutral pH as an inactive zymogen. Upon acidification, it is activated into its mature form through a series of intramolecular processing events. High-resolution X-ray crystal structures for both an active low pH, (pH 5.1) and an inactive high pH (pH 7.5) forms have been determined in an effort to design selective inhibitors for CatD (Lee et al., 1998). Based on the crystal structure, the hydroxyl group of tyrosine 10 (Y10) forms a hydrogen bond with the ionized catalytic carboxylate residue of aspartate 33 (Asp33), thus stabilizing the inactive form. We hypothesize that changing Y10 to phenylalanine (F) should favor the open conformation of our enzyme since phenylalanine cannot hydrogen bond with the catalytic carboxylate of Asp33. Site-directed mutagenesis was performed on “short” recombinant pseudocathepsin D to create the Y10F variant. The enzyme was purified by affinity chromatography followed by an ion exchange chromatography yielding purified mature enzyme. Additionally, refolding and pH experiments were performed with wild type and variants in order to optimize the purification yield. [Supported by NIH grant DK18865]

INTRODUCTION

Cathepsin D (CatD) is a member of the pepsin family of aspartic peptidases. Aspartic peptidases are endopeptidases, consisting of two aspartic acid residues that catalyze peptide bond hydrolysis through a general base catalyzed mechanism. A low pH optimum for catalytic activity is an important characteristic for these lysosomal enzymes. However, it has also been detected in other places including the cytosol and the extracellular space. In the extracellular space where pH is more neutral, it has been implicated in pathological processes such as inflammation, tumor progression, formation of metastases, and apoptosis (Leto et al., 1992; Mignatti and Rifkin, 1993). These findings are interesting in light of the acidic pH optimum of Cathepsin D.

To gain more insight on the role of CatD at more neutral pH, Lee and colleagues solved the structure at a high pH. They proposed a reversible, pH-dependent conformational change from a low pH (CatDlo), “open” and active conformation, to a high pH (CatDhi), “closed”, inactive conformation (1998). Structural comparison of CatDhi and CatDlo reveal significant conformational differences. An increase in pH induces a set of conformational changes that results in the relocation of the N-terminal segment into the active site where both catalytic aspartates are likely ionized. Additionally, the positively charged Nitrogen atom of Lysine 8 (K8) along with the hydrogen bond formed with the hydroxyl group of Y10 form interactions with the ionized catalytic aspartate acid residues to stabilize the inactive form. Reactivation due to a decrease in pH can occur due to protonation of one of the two catalytic aspartates, thus weakening electrostatic interactions and hydrogen bonds between the catalytic aspartates and Y10, respectively.

The goal of this project was to test the significance of hydrogen bonding between Y10 and Asp 33 in stabilizing the inactive form of the enzyme. The recombinant model, “short” pseudocathepsin D (rHuCatD) used to test our hypothesis is kinetically similar to the mature enzyme (Beyer and Dunn, 1996). By mutating the Y10 residue in “short” rHuCatD to phenylalanine (F), we expect the variant Y10F to exhibit an open conformation due to its inability to hydrogen bond with the catalytic carboxylate of Asp33.

MATERIALS AND METHODS

Mutagenesis

The gene encoding enzyme “short” rHuCatD obtained from Brian Beyer (1996) was mutated using the Quik-changeTM Site Directed Mutagenesis Kit from Stratagene (La Jolla, CA) to make the variant, Y10F. Shown in Table 1 is the primer and its reverse complement used to introduce the point mutation.

Table 1
Primer and the Reverse Complement
Primer Sequence Desired Mutation
5' CCCATTCCCGAGGTGCTCAAGAACTTCATGGACC 3' Tyr10Phc
3' GGGTAAGGGCTCCACGAGTTCTTGAAGTACCTG 5' Tyr10Phc


Following the PCR reaction, the entire region encoding rHuCatD was sequenced at the University of Florida ICBR DNA Sequencing Core Laboratory to confirm the presence of the intended mutation. Constructs bearing the mutation were then transformed into E. coli BL21(DE3)pLysS cells for overexpression (Stratagene) according to manufacturer’s protocol. The transformation mixture was plated on LB agar plates containing 50 μg/mL ampicillin for 15-17 hours. Isolated colonies were selected and cultured overnight in 5 mL LB media containing ampicillin (50 μg/mL). Double stranded plasmid DNA was then purified using Quiagen miniprep plasmid purification columns.

Expression

The protocol of Chen et al. (1991) was modified as follows in order to express and purify the variant. A 1:50 dilution of an overnight culture grown in M9 media (10 mg/mL thiamine, 0.5% casamino acids, 0.2% glucose) containing 50 mg/L ampicillin was made into LB media containing 50 mg/L ampicillin. As soon as the mixture was grown to an OD600 of 0.5, IPTG was added to give a final concentration of 0.5 mM. Aliquots were taken before adding IPTG at 0, 1, 2, and 3 hours intervals, and were analyzed by SDS-PAGE and Coomassie Blue staining in order to observe a time-dependent increase in expression. The cells were pelleted by centrifugation and resuspended in 4.2 mLs of 50 mM Tris-HCl pH 7.4, 150 mM NaCl and 1 mM MgCl2 (buffer A) per gram of cells. Following the addition of 80 Kunitz units of Dnase per mL of suspension, the cells were lysed twice through a French Pressure Cell. The resulting lysate was layered over 27% sucrose and centrifuged in order to isolate inclusion bodies. The inclusion bodies were resuspended in buffer A containing 1% Triton X-100 and pelleted again through the sucrose cushion. The pellet weight was measured and the inclusion bodies were frozen at –80°C.

Resolubilization of Inclusion Bodies

To recover the active enzyme, the inclusion bodies were resolubilized by rapid dilution following previously reported procedures (Conner and Richo, 1992; Scarborough et al., 1993; Beyer and Dunn, 1996). Inclusion bodies were solubilized and reduced at a concentration of 2 mg/mL in 50 mM CAPS pH 10.7, 50 mM _-mercaptoethanol, and 8 M urea. The resolubilized solution was stirred for 30 minutes at 25°C and was diluted 100 fold by dropwise addition into filtered 10 mM Tris-HCl pH 8.7 to a final concentration of 20 µg/mL. After 4 hours, oxidized glutathione was added to the refolding solution and the variant was allowed to refold for a pre-determined optimum refolding time. The refolded material was then activated by acidification with formic acid to pH 3.7. Refolding time-course and activation-time course experiments were performed by removing aliquots from the refolding and activated material respectively, and monitoring for emergence of enzymatic activity.

Purification

The activated protein solution was applied to a C 16/20 affinity column (Amersham-Pharmacia) containing 1 mL of resuspended pepstatinyl-agarose (Huang et al., 1979). The column was then washed with at least 5 mLs of 0.01 M sodium formate pH 3.7, 0.4 M NaCl, 0.05% Brij-35. It was eluted with cold 20 mM Tris-HCl pH 8.0, 0.4 M NaCl and 0.5% Brij 35 and fractions were analyzed for catalytic activity. The fractions exhibiting activity were pooled and dialyzed against 4 L of 10 mM Tris-HCl pH 8.0 overnight at 4°C to desalt. Ion-exchange chromatography was used to purify the mature form of the enzyme. A 1 mL HiTrap Q Sepharose HP (Amersham Pharmacia), an anion exchanger, was equilibrated with 10 mM Tris-HCl pH 8.0 (buffer A). The desalted material was then applied to the column at a flow rate of 1 mL/min. The column was washed with buffer A and the mature protein was eluted with a salt gradient of 0-60 mM.

Spectrofluorometric analysis

Enzyme catalyzed hydrolysis of a fluorogenic substrate was spectrofluorometrically monitored on a PerSeptive Biosystems CytoFluor Series 4000/TC Multi-Well Plate Reader. Samples were excited at 360 nm and emission was monitored at 460 nm.

pH-dependent stability

Enzyme was preincubated for 10 minutes at 37°C in the following 100 mM buffers: glycine-HCl pH 2.0-3.0; sodium formate pH 3.75; sodium acetate pH 4.5; MES pH 5.5-7.0; Tris-HCl pH 8.3-9.3; CAPS pH 10.7-11.7. After 10 minutes, pH was recorded and the enzyme was added to 0.2 M sodium formate pH 3.7 at 37°C for an additional 3 minutes. It was then added to 30 _M substrate and enzymatic activity was monitored. The final pH was recorded for all points.

RESULTS

Purification of Cat D and variants was achieved by expression in E. coli, however, the protein was deposited into insoluble inclusion bodies. Approximately 1 gram weight of inclusion body pellet was harvested from a 1 L over-expression. SDS-PAGE analysis showed a time-dependent increase in expression of the target protein (~40kDa) after addition of IPTG (Figure 1). Since the enzyme was synthesized as an inactive proenzyme, upon acidification and after a series of purification steps, the mature enzyme was obtained (Figure 4). First, affinity chromatography was used to purify the variant from the activated refolding mixture (Figure 2). An ion-exchange chromatography was included to purify the mature form of the enzyme from the post affinity purification (Figure 3).
Experiments to determine the optimal refolding time and activation time for each variant were performed (Figure 5, Figure 6, Table 1). These experiments indicated that the optimal refolding and activation time varies with the variant. Moreover, addition of 20% glycerol to the refolding mixture did not yield any significant improvements in refolding process (Figure 10).

Comparing the activity of optimally refolded and activated WT and variants, it is apparent that the variants refold and/or activate less efficiently compared to the WT material (Figure 7). Experiments testing the effect of pH on refolding indicated that refolding at pH 10.5 and 11 increased the amount of recovered active protein for D187R and E5QE180Q, respectively (Figure 8). Additionally, the pH stability assay for Y10F was performed to show the stability of the variant over a broad pH range compared to wild type. The assay shows Y10F exhibited significant pH stability between pH 3.0 and pH 6.0 (Figure 9).

Figure 1. SDS-PAGE analysis showing a time-dependent increase in expression Y10F (~40kDa) after addition of IPTG


Figure 1. SDS-PAGE analysis showing a time-dependent increase in expression Y10F (~40kDa) after addition of IPTG. Left lane corresponds to molecular weight markers. Lane #1 shows E. coli proteins before induction. Lanes #2-4 illustrate over-expression of Y10F. Lane 5 corresponds to the proteins present in the supernatant. Lane #6 shows the purity of the inclusion body preparation.

Figure 2. Elution profile of Y10F Post pepstatinly agarose
Figure 2. Elution profile of Y10F Post pepstatinly agarose. Activity measured by monitoring the cleavage of fluorogenic substrate. Purified enzyme was pooled from fractions 1 to 6.

Figure 3. Elution profile of Y10F for DEAE ion exchange chromatography
Figure 3. Elution profile of Y10F for DEAE ion exchange chromatography. Activity measured by monitoring the cleavage of fluorogenic substrate. A salt gradient of 0-60 mM was determined to achieve separation of the mature enzyme.

Figure 4. Silver stained SDS-PAGE depicting purification of Y10F.

Figure 4. Silver stained SDS-PAGE depicting purification of Y10F. The far left lane corresponds to molecular weight markers. Lane #3 represents full length Y10F from the refolding mixture. Lane #2 shows the pepstatinly-agarose purification. Lane #1 illustrates the DEAE ion-exchange for Y10F yielding the mature form of the enzyme.

Figure 5. Refolding time-course experiment for recombinant “short” pseudocathepsin D and its variants.

Figure 5. Refolding time-course experiment for recombinant “short” pseudocathepsin D and its variants.

Figure 6. Activation time-course experiment for recombinant “short” pseudocathepsin D and its variants.

Figure 6. Activation time-course experiment for recombinant “short” pseudocathepsin D and its variants.

Figure 7. Activity of WT and variants after optimal refolding and activation.

Figure 7. Activity of WT and variants after optimal refolding and activation.

Figure 8. pH dependent refolding for D187R (red), E180Q (green), and E5QE180Q (black).

Figure 8. pH dependent refolding for D187R (red), E180Q (green), and E5QE180Q (black). Refolding of D187R was at pH 10.5, E180Q at pH 8.7 and E5QE180Q at pH 11.0.

Figure 9. pH-dependent stability assay for Y10F and WT

Figure 9. pH-dependent stability assay for Y10F and WT. WT is plotted as closed squares; Y10F open squares. Activity measured from cleavage of fluorogenic substrate.

Figure 10. Refolding time course assay for WT showing the effect of addition of 20% glycerol.

Figure 10. Refolding time course assay for WT showing the effect of addition of 20% glycerol.

Table 1
Optimum refolding and activation time for WT and variants. The pH at which each enzyme was refolded and activated is also shown
Variant Optimal Refolding Time (days) pH of refolding reaction Optimal Activation Time (days) Activation pH
WT 3 8.7 1 3.7
E180Q 1 8.7 1 3.7
E5QE180Q 1 11.0 2 3.7
D187R 0.5 10.5 2 3.7
Y10F 2 8.7 3 3.7
Y10FE180Q 1 8.7 2 3.7

CONCLUSIONS

Since the refolding step was very inefficient with about a 1% recovery of the active material (Beyer and Dunn, 1996), parameters such as amount of affinity resin and flow rate of affinity chromatography were tested. Additionally, glycerol was added and pH of the refolding mixture was varied in order to optimize the refolding process.

This study allowed us to determine optimal refolding conditions for Y10F and other variants. Our data showed substantial decrease in the amount of recovered active material compared to WT. Refolding and activation time-course experiments enabled us to optimize the purification yield for WT and variant enzymes. The data obtained from this study provided valuable information that enabled further kinetic studies.

Sophisticated experiments such as pH-dependent Michaelis-Menten kinetics, intrinsic tryptophan fluorescence experiments, and N-terminal sequencing for Y10F were performed by Nathan Goldfarb in Dr. Dunn’s laboratory at the University of Florida. These studies revealed that Y10F, in fact exhibited a conformational equilibrium shift to the more open, “active” form of the enzyme. This indicated that the hydrogen bonding interaction between Y10 and Asp 33 is a crucial interaction that stabilizes the closed, high pH form of the enzyme. However, additional studies need to be done to address the role of the Lys 8 - Asp 33 interaction in this pH dependent conformational change.

REFERENCES

Beyer BM, Dunn BM. 1996. Self-activation of recombinant human lysosomal procathepsin D at a newly engineered cleavage junction, "short" pseudocathepsin D. J Biol Chem 271:15590-15596.

Chen Z, Koelsch G, Han HP, Wang XJ, Lin XL, Hartsuck JA, Tang J. 1991. Recombinant rhizopuspepsinogen. Expression, purification, and activation properties of recombinant rhizopuspepsinogens. J Biol Chem 266:11718-11725.

Conner GE, Richo G. 1992. Isolation and characterization of a stable activation intermediate of the lysosomal aspartyl protease cathepsin D. Biochemistry 31:1142-1147.

Huang JS, Huang SS, Tang J. 1979. Cathepsin D isozymes from porcine spleens. Large scale purification and polypeptide chain arrangements. J Biol Chem 254:11405-11417.

Lee AY, Gulnik SV, Erickson JW. 1998. Conformational switching in an aspartic proteinase. Nat Struct Biol 5:866-871.

Leto G, Gebbia N, Rausa L, Tumminello FM. 1992. Cathepsin D in the malignant progression of neoplastic diseases (review). Anticancer Res 12:235-240.

Mignatti P, Rifkin DB. 1993. Biology and biochemistry of proteinases in tumor invasion. Physiol Rev 73:161-195.

Scarborough PE, Guruprasad K, Topham C, Richo GR, Conner GE, Blundell TL, Dunn BM. 1993. Exploration of subsite binding specificity of human cathepsin D through kinetics and rule-based molecular modeling. Protein Sci 2:264-276.

--top--

Back to the Journal of Undergraduate Research