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Published online before print May 2, 2006, 10.1110/ps.062129506
Protein Science (2006), 15:1277-1289. Published by Cold Spring Harbor Laboratory Press. Copyright © 2006 The Protein Society
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Contribution of the mu loop to the structure and function of rat glutathione transferase M1-1

Jennifer L. Hearne and Roberta F. Colman

Department of Chemistry and Biochemistry, University of Delaware, Newark, Delaware 19716, USA

(RECEIVED February 1, 2006; FINAL REVISION February 24, 2006; ACCEPTED February 24, 2006)


   Abstract
TOP
 Abstract
Introduction
 Results
 Discussion
 Materials and methods
 References
 
The "mu loop," an 11-residue loop spanning amino acid residues 33–43, is a characteristic structural feature of the mu class of glutathione transferases. To assess the contribution of the mu loop to the structure and function of rat GST M1-1, amino acid residues 35–44 (35GDAPDYDRSQ44) were excised by deletion mutagenesis, resulting in the "Deletion Enzyme." Kinetic studies reveal that the Km values of the Deletion Enzyme are markedly increased compared with those of the wild-type enzyme: 32-fold for 1-chloro-2,4-dinitrobenzene, 99-fold for glutathione, and 880-fold for monobromobimane, while the Vmax value for each substrate is increased only modestly. Results from experiments probing the structure of the Deletion Enzyme, in comparison with that of the wild-type enzyme, suggest that the secondary and quaternary structures have not been appreciably perturbed. Thermostability studies indicate that the Deletion Enzyme is as stable as the wild-type enzyme at 4°C and 10°C, but it rapidly loses activity at 25°C, unlike the wild-type enzyme. In the temperature range of 4°C through 25°C, the loss of activity of the Deletion Enzyme is not the result of a change in its structure, as determined by circular dichroism spectroscopy and sedimentation equilibrium centrifugation. Collectively, these results indicate that the mu loop is not essential for GST M1-1 to maintain its structure nor is it required for the enzyme to retain some catalytic activity. However, it is an important determinant of the enzyme's affinity for its substrates.

Keywords: glutathione S-transferase M1-1; deletion mutagenesis; mu loop; substrate specificity


   Introduction
TOP
 Abstract
 Introduction
Results
 Discussion
 Materials and methods
 References
 
Glutathione transferases (GSTs, E.C. 2.5.1.18 [EC] ) constitute a family of enzymes involved in Phase II detoxification of endogenous and exogenous xenobiotics (Jakoby and Habig 1980). GSTs are ubiquitous but are particularly abundant in hepatic tissue, comprising ~4% of the cytosolic protein content in hepatocytes (Eaton and Bammler 1999). These enzymes detoxify xenobiotics by catalyzing the nucleophilic addition of the thiolate of glutathione (GSH) to substrates bearing an electrophilic center. The conjugate is rendered more water-soluble than the original xenobiotic, which is advantageous for excretion, usually as part of the mercapturic pathway (Mannervik and Danielson 1988; Boyer 1989; Pickett and Lu 1989; Coles and Ketterer 1990; Armstrong 1991). Physiologically, GSTs have been implicated in the protection against carcinogenesis, as well as in drug resistance (Soberman and Austen 1989; Waxman 1990).

The cytosolic GSTs are subdivided into at least seven classes based on their sequence identity, physical properties, and substrate and inhibitor specificities (Mannervik et al. 1985; Meyer et al. 1991; Pemble et al. 1996; Rossjohn et al. 1998; Board et al. 2000; Pettigrew and Colman 2001; Hayes et al. 2005). Isozymes within a class exhibit at least 40% sequence identity, while the identity between classes is less than 25%, although the tertiary structure is generally conserved (Sheehan et al. 2001; Hayes et al. 2005). Table 1 shows a sequence alignment of the first 58 amino acid residues of rat GST M1-1 with the Deletion Enzyme, rat GST A1-1, and human GST P1-1. Despite the low sequence identity among the classes of GSTs, there are several features shared among the most prevalent GSTs (alpha, mu, and pi classes). These classes have a similar fold in which the subunit structure is divided into two domains (Armstrong 1997). The N-terminal domain is the more highly conserved domain; it functions in providing most of the binding contacts for GSH (Armstrong 1997). Its topology is similar to that of thioredoxin (beta{alpha}beta{alpha}betabeta{alpha}) but lacks the Cys-X-X-Cys motif (Martin 1995; Murzin et al. 1995). The C-terminal domain is responsible for providing most of the contacts in the xenobiotic substrate-binding site (Armstrong 1997). This fold consists of five {alpha}-helices in the case of the mu and pi classes and six in the alpha class (Sheehan et al. 2001). The overlapping but distinct substrate and inhibitor specificities between classes may be due to the sequence differences in the C-terminal domain (Wilce and Parker 1994). Although there are differences in subunit–subunit rotation and active site solvent exposure, the active site of each class of GSTs is in an equivalent position in the enzyme structure (Sinning et al. 1993).


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  		Table 1. A partial sequence alignment of GST M1-1, the Deletion Enzyme, GST A1-1, and GST P1-1

 
There are characteristic secondary structural features for the alpha and mu classes: The alpha class has an extra amphipathic C-terminal helix ({alpha}-helix 9), which packs onto the hydrophobic substrate site, resulting in a more hydrophobic, as well as a smaller, substrate site than in the mu and pi classes (Sinning et al. 1993; Dirr and Wallace 1999; Sheehan et al. 2001). This extra helix functions in ligandin binding and catalytic activity but contributes minimally to the overall stability of the enzyme structure (Dirr and Wallace 1999). As shown highlighted in red in Figure 1A (dimeric GST M1-1) and 1B (monomeric GST M1), the distinctive feature of the mu class is the mu loop, which is located between beta-strand 2 and {alpha}-helix 2 and spans amino acid residues 33–43 (Ji et al. 1992) in the highly conserved N-terminal domain, adjacent to the xenobiotic substrate site. It is one of three structural elements that creates an active site cleft which is more constricted and less exposed to solvent than in the pi class (Wilce and Parker 1994).


Figure 1
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  		Figure 1. (A) Structure of the dimeric form of GST M1-1 (PDB code 6GST; Xiao et al. 1996). (Cyan) Subunit A; (white) subunit B. The mu loop of each subunit, amino acid residues 33–43 as well as the amino acid residue 44, is highlighted in red. (B) Structure of one subunit of the dimer of GST M1-1 (PDB code 6GST). The mu loop, amino acid residues 33–43, is highlighted in red. GSH is shown as it was crystallized with the wild-type enzyme; GSH is colored in green. Yellow arrows show the approximate location of the alpha carbon of amino acid residues 35 and 44. The monomer has been rotated 90° around the Y-axis to provide a better view of the mu loop.

 
In this study, we investigated the contribution of the mu loop, generally considered to include residues 33–43, to the structure and function of GST M1-1. Using deletion mutagenesis, amino acid residues 35–44 of the mu loop were excised. These amino acid residues were chosen for deletion because in silico modeling of the Deletion Enzyme, in comparison with the wild-type enzyme, predicted that the deletion of these amino acid residues would not appreciably impact the structure of the protein backbone. The excision of these amino acid residues results in a new peptide bond between Met34 and Trp45, which are relatively close in the wild-type enzyme structure (Fig. 1). This linkage does not introduce a repulsive interaction. (Other amino acid residue combinations were considered for excision, such as 35–41 and 33–44, but were not explored further because of deviations in the protein backbone structures as revealed by in silico modeling.) We now address the following questions: Does the mu loop provide stability to the structure of the wild-type enzyme? and, is the mu loop required for normal enzyme catalysis?


   Results
TOP
 Abstract
 Introduction
 Results
Discussion
 Materials and methods
 References
 
Enzyme purity and characterization of the primary structure of the Deletion Enzyme
The plasmid encoding the wild-type enzyme rat (Rattus norvegicus) GST M1-1 or the Deletion Enzyme was expressed in JM105 Escherichia coli cells and purified using chromatography on S-hexylglutathione agarose, as discussed in Materials and Methods. Typically, 15 mg/L cell culture of the wild-type GST M1-1 enzyme and 8 mg/L cell culture of the Deletion Enzyme were obtained. N-terminal sequencing (15 reaction cycles) indicated, in both cases, the presence of only one protein with the phenylthiohydantoin amino acid derivative sequence, PMILGYWNVRGL, which is unique to rat GST M1-1, establishing the purity of each enzyme. N-terminal sequencing for 47 reaction cycles confirmed that residues 35–44 were absent in the Deletion Enzyme (data not shown).

The theoretical monomer molecular weight of the wild-type enzyme and the Deletion Enzyme was calculated using Compute pI/Mw on the Expasy website (www.expasy.ch), as 25,783 g/mol and 24,677 g/mol respectively, a difference of 1105 g/mol. Electrospray ionization mass spectrometry (ESI-MS) results reveal one predominant peak for each enzyme sample with an average molecular mass of 25,827 for the wild-type enzyme monomer (monomer plus two sodium atoms) and a molecular mass of 24,677 for the Deletion Enzyme monomer (data not shown). These results indicate that the enzymes are pure and of the expected molecular mass. The Deletion Enzyme's monomer molecular mass also affirms the deletion of the 10 residues.

Subunit interaction
The crystal structure of GST M1-1 (PDB code 6GST; Xiao et al. 1996) shows a dimeric enzyme. In this study, sedimentation equilibrium experiments conducted at 4°C indicate the weight average molecular weights of the wild-type enzyme and the Deletion Enzyme at 0.3 mg/mL are 45.8 ± 0.1 kDa and 44.2 ± 0.1 kDa, respectively. To find out if glutathione (GSH) aids in dimerization of either enzyme, GSH was added to the enzyme samples and the weight average molecular weights were determined. In the presence of 2.5 mM GSH, the wild-type enzyme exhibits a weight average molecular weight of 48.0 ± 0.1 kDa and the Deletion Enzyme exhibits a weight average molecular weight of 46.9 ± 0.1 kDa in the presence of 5 mM GSH. Similar results were obtained for enzyme samples at 0.06 mg/mL in the absence and presence of GSH. These results indicate that both the wild-type enzyme and the Deletion Enzyme are present predominantly in the dimeric form in solution, demonstrating that the normal subunit interaction has not been affected by the deletion of amino acid residues 35–44.

Conformation of the wild-type enzyme and the Deletion Enzyme
To determine if the Deletion Enzyme's secondary structure is similar to that of the wild-type enzyme, the circular dichroism (CD) spectrum of each enzyme was recorded at 4°C (data not shown). The spectrum of the Deletion Enzyme deviates somewhat from that of the wild-type enzyme; it exhibits a smaller negative molar ellipticity between 208 nm and 215 nm, indicating a loss of {alpha}-helical structure; however, the molar ellipticity at 222 nm of the Deletion Enzyme is similar to that of the wild-type enzyme. These results indicate that the deletion of amino acid residues 35–44 does not greatly affect the secondary structure of the enzyme.

Fluorescence spectroscopy can be used to probe the environment of tyrosine, phenylalanine, and tryptophan residues in an enzyme (Lakowicz 1983; Luo et al. 2002). The emission maximum of tryptophan is often the result of the sensitivity of the indole ring to the solvent polarity. Therefore, the blue shift in the maximum emission of the wild-type enzyme and the Deletion Enzyme compared with that of free tryptophan is most likely due to shielding of the tryptophan residues in the proteins from the aqueous phase (Lakowicz 1983). In Figure 2, a comparison of the emission spectrum (excitation at 280 nm) of the wild-type enzyme and free tryptophan (at the same concentration as in the enzyme subunit) reveals that the spectrum of the wild-type enzyme is mainly due to the contribution of the four tryptophan amino acid residues per subunit (Trp7, Trp45, Trp146, and Trp214), despite tyrosine's high absorbance and quantum yield at 280 nm. These results are not unusual: As is generally the case, the fluorescence of tryptophan dominates the spectrum and tyrosine remains undetected (Lakowicz 1983). The wild-type enzyme has an emission maximum (330 nm) blue-shifted 18 nm, as compared with that exhibited by free tryptophan (348 nm), indicating that the tryptophan residues in the enzyme are shielded from the aqueous phase by the enzyme.


Figure 2
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  		Figure 2. The steady-state fluorescence spectra of the wild-type enzyme and the Deletion Enzyme (2.2 x 10–5 M enzyme subunits) in 0.1 M potassium phosphate buffer (pH 6.5) containing 1 mM EDTA at 4°C were measured on a Perkin Elmer MPF-3 fluorescence spectrophotometer. Emission spectrum (excitation at 280 nm, bandwidth of 10 nm) of GST M1-1 (bullet), the Deletion Enzyme ({blacktriangledown}), and free tryptophan ({blacksquare}).

 
A comparison of the emission spectrum (excitation at 280 nm) of wild-type enzyme and the Deletion Enzyme reveals that, under these conditions, the wild-type enzyme's relative fluorescence intensity is 25 at its emission maximum, while the Deletion Enzyme exhibits 156% relative fluorescence intensity at its emission maximum (333 nm) compared with that of the wild-type enzyme. The emission maximum shift of the Deletion Enzyme is likely due to the change in solvent exposure of Trp35 in the Deletion Enzyme (equivalent to Trp45 in the wild-type enzyme) (Lakowicz 1983). The increase in fluorescence intensity of the Deletion Enzyme compared with that of the wild-type enzyme may be due to the solvent exposure of Trp35. Hornby et al. (2000) demonstrated that the unfolding of GST M1-1 caused an increase in the fluorescence intensity exhibited by the enzyme. Although we did not unfold the Deletion Enzyme, the structure of the enzyme renders Trp35 solvent accessible, presumably the cause of the increase in fluorescence intensity. The homology model of the Deletion Enzyme supports this explanation (see Discussion).

Thermostability experiments were conducted to explore whether the mu loop has a structural function in the wild-type enzyme. The wild-type enzyme (0.3 mg/mL) and the Deletion Enzyme (0.3 mg/mL) were incubated at 4°C, 10°C, or 25°C and at various times were tested for catalytic activity at 25°C using 1-chloro-2,4-dinitrobenzene (CDNB) and GSH as substrates. The results are shown in Figure 3. The wild-type enzyme is stable at each temperature tested for the duration of the experiment (60 min). Figure 3A shows the wild-type enzyme's stability at 4°C. As shown in Figure 3A,B the Deletion Enzyme is as stable as the wild-type enzyme at 4°C and 10°C. However, incubation of the Deletion Enzyme at 25°C results in the loss of ~50% of its activity in about 15 min (Fig. 3C).


Figure 3
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  		Figure 3. Thermostability study of the wild-type enzyme and the Deletion Enzyme at 4°C, 10°C, and 25°C. In each panel, wild-type enzyme is represented by the black triangles ({blacktriangleup}) connected by a dashed line and the Deletion Enzyme is represented by the black circles (bullet) connected by a solid line. The rates of inactivation of the wild-type enzyme and the Deletion Enzyme were monitored at 4°C (A) 10°C (B) and 25°C (C). The activity of the enzymes is expressed as Et/Eo (observed activity/initial activity).

 
Both the wild-type enzyme and the Deletion Enzyme (0.06 mg/mL and 0.3 mg/mL) were also monitored over 76 h at 4°C. Enzyme samples at 0.3 mg/mL were determined to be stable over the time period necessary to conduct the sedimentation equilibrium experiments (data not shown). At 0.06 mg/mL the wild-type enzyme was stable for the 76-h incubation but the Deletion Enzyme lost ~85% of its activity. It is also apparent that the loss of activity of the Deletion Enzyme is not due to a change in the quaternary structure since the same weight average molecular weight was determined and found to be unchanged.

When the wild-type enzyme and the Deletion Enzyme (both at 0.15 mg/mL) were monitored as a function of temperature in the range of 4°C–25°C, the circular dichroism spectra did not change. The CD spectrum of the Deletion Enzyme at 25°C was recorded after 15 min of incubation (a time period giving ~50% loss of activity). Therefore, the perturbation of activity of the Deletion Enzyme compared with that of the wild-type enzyme is not due to an appreciable change in the enzyme's secondary structure.

Kinetic parameters of the wild-type enzyme and the Deletion Enzyme
For all kinetic measurements it was imperative to maintain the Deletion Enzyme at 4°C until immediately before the measurement which was taken at 25°C. The activities of the wild-type enzyme and the Deletion Enzyme were compared by measuring the enzymes' abilities to catalyze the reaction of GSH with two xenobiotic substrates, CDNB and monobromobimane (mBBr) at pH 6.5. Table 2 lists the kinetic parameters of the wild-type enzyme and the Deletion Enzyme for CDNB (data columns 1–3) and mBBr (data columns 6 and 7). The Km values of the Deletion Enzyme drastically increase: 32-fold for CDNB (column 2), 99-fold for GSH (column 3), and 880-fold for mBBr (column 7). The Vmax value for each substrate of the Deletion Enzyme increases modestly: VmaxCDNB increases approximately threefold (column 1) and VmaxmBBr increases approximately eightfold (column 6). The difference between the effects of the deletion on the enzyme, as seen in the kinetic parameters, for the two xenobiotic substrates, is consistent with our previous finding that CDNB and mBBr occupy distinct sites (Hearne and Colman 2005). The results demonstrate that the excision of amino acid residues 35-44 appreciably perturbs the enzyme's affinity for the xenobiotic substrates. Wild-type rat GST A1-1 and human GST P1-1 kinetic parameter values are included in Table 2 for comparison.


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  		Table 2. A comparison of the CDNB, GSH, mBBr kinetic parameters of GST M1-1, GST A1-1, GST P1-1, and the Deletion Enzyme

 
To determine if the effect of removing amino acid residues 35–44 was localized to the glycyl domain of GSH, glutathione monoethyl ester (glycyl carboxylate esterified, GSHME) was tested as the nucleophilic substrate in the CDNB reaction. Table 2 shows that the Vmax value (cf. column 4 with column 1) of the wild-type enzyme is unchanged and the Km is decreased twofold (cf. column 5 with column 3) in comparison with these values obtained for GSH. The effect on Km of substituting GSHME for GSH as the nucleophilic substrate in the CDNB reaction is the same for the Deletion Enzyme as it is for the wild-type enzyme (cf. column 5 with column 3). In contrast, we find that GSHME decreases the Vmax value of the Deletion Enzyme fivefold (cf. column 4 with column 1) in comparison with the Vmax value attained when GSH is the nucleophilic substrate in the CDNB reaction. Collectively, these results suggest that the effect of deleting amino acid residues 35–44 is not confined to the glycyl domain of the GSH binding site. The decreased affinity of the Deletion Enzyme for GSH and its derivative is most likely a net effect of changing the hydrogen bonding network and electrostatic interactions within the GSH site.

A major function of the glutathione transferases is to lower the pKa of the thiol of GSH from 9.13, characteristic of GSH in solution, to ~6, which allows the enzyme-bound glutathione to exist as the thiolate at physiological pH and, therefore, to be a more effective participant in the nucleophilic aromatic substitution reaction (Wang et al. 1996). To probe the effect of the elimination of amino acid residues 35–44 on the pKa of the enzyme-bound GSH, the kcat and the KmCDNB of the Deletion Enzyme using CDNB and GSH as substrates were determined over the pH range of 5.0–7.0 and compared with those values of the wild-type enzyme in the pH range of 5.75–8.0, with the results shown in Figure 4. It was determined that both enzymes are stable in the pH range used in this evaluation (data not shown). An evaluation of kcat/KmCDNB values reveals that the Deletion Enzyme is ~10 times less efficient in catalysis than the wild-type enzyme at pH 6.5 (Deletion Enzyme = 50,420 sec–1 µM–1 and wild-type enzyme = 497,567 sec–1 µM–1). However, as the pH is decreased, the kcat/KmCDNB versus pH curves of the two enzymes come closer together, and at pH 5.75 the Deletion Enzyme is only five times less efficient in catalysis than the wild-type enzyme. These results indicate that the pKa of the enzyme–glutathione complex is decreased in the Deletion Enzyme (Wang et al. 1996). Analysis of kcat/KmCDNB versus pH reveals that the pKa for the Deletion Enzyme–GSH complex is 5.4 ± 0.1, 0.5 pH units lower than the pKa of the wild-type enzyme (5.9 ± 0.1; Fig. 4). These results show that, although the affinity for GSH is weakened in the Deletion Enzyme, the enzyme retains its ability to lower the pKa of the enzyme-bound GSH; in fact, the pKa is lower than that of the wild-type enzyme. Therefore, the altered pH dependence cannot account for the lowered kcat/KmCDNB of the Deletion Enzyme.


Figure 4
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  		Figure 4. pH dependence of kcat/KmCDNB for the wild-type enzyme (A) and the Deletion Enzyme (B). These values were determined by fitting the data to the equation pKa = (kcat/KmCDNB)/(1 + 10(pKa–pH)), where kcat is the turnover number of the enzyme for the substrates GSH and CDNB, the Km is that of CDNB, and the pH is that at which the catalytic measurements were recorded.

 

   Discussion
TOP
 Abstract
 Introduction
 Results
 Discussion
Materials and methods
 References
 
This study demonstrates that the mu loop of GST M1-1 is not required for the enzyme's catalytic function nor is it a major determinant of the secondary or quaternary structure of the enzyme. However, it is important in the enzyme's affinity for the substrates tested.

It is not surprising to find that the deletion of amino acid residues 35–44 has little effect on the weight average molecular weight (or extent of association of the subunits) of the enzyme. Figure 1 clearly shows that the mu loop is located far from the subunit–subunit interface. The homology model of the Deletion Enzyme was based on the structure of subunit B of GST M1-1 entries in the protein data bank (PDB). An overlay of the wild-type monomer (PDB code 6GST; Xiao et al. 1996) and the Deletion Enzyme monomer is shown in Figure 5A. There is excellent correspondence between the structures of the enzymes except in the immediate vicinity of the mu loop. The similarity between the wild-type enzyme structure and that of the homology model of the Deletion Enzyme is in agreement with our experimental results indicating that the Deletion Enzyme's secondary structure is not appreciably perturbed.


Figure 5
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  		Figure 5. (A) Overlay of GST M1-1 (6GST, white) and the homology model of the Deletion Enzyme (blue) showing the difference in solvent exposure of Trp45 (GST M1-1, red) and Trp35 (Deletion Enzyme, yellow). The mu loop of the wild-type enzyme is shown in red and the remaining mu loop region of the Deletion Enzyme is shown in yellow. Please note that amino acid residue 45 of wild-type enzyme is equivalent to amino acid residue 35 of the Deletion Enzyme. (In the wild-type enzyme residues 45 to 217 are equivalent to those numbered 10 less in the Deletion Enzyme. Specifically, Trp45 of the wild-type enzyme is equivalent to Trp35 of the Deletion Enzyme. The other tryptophan residues present in each enzyme are also displayed. GSH is shown as bound in wild-type enzyme and is colored green. (B) Enlargement of the active site region of the wild-type enzyme and the Deletion Enzyme as shown in A. The coloring remains the same with the exception that the GSH molecule is colored by atom (oxygen, red; nitrogen, blue; sulfur, yellow; carbon, white).

 
It may be that there are changes in orientation of amino acid residues in the vicinity of the remaining mu loop and the active site in the Deletion Enzyme that do not appreciably impact the secondary structure but could affect the binding of substrates. Again, the homology model of the Deletion Enzyme proved useful in understanding these changes. Figure 5A shows the in silico predicted change in environment of Trp35 of the Deletion Enzyme (equivalent to Trp45 in the wild-type enzyme). (In the wild-type enzyme, residues 45 to 217 are equivalent to those numbered 10 less in the Deletion Enzyme. Specifically, Trp45 of the wild-type enzyme is equivalent to Trp35 of the Deletion Enzyme.) In contrast to Trp35, the three other tryptophan amino acid residues are not appreciably displaced. This predicted change in environment of Trp35 of the Deletion Enzyme is consistent with the gain in the relative fluorescence intensity and the red shift in maximum wavelength of the Deletion Enzyme compared with that of the wild-type enzyme.

The catalytic role of the mu loop was investigated by comparing the kinetic parameters of GST A1-1, M1-1, and P1-1 isozymes with the Deletion Enzyme for the reaction of GSH with two characterized xenobiotic substrates, CDNB and mBBr. The mu loop and the folding of the C terminus into a cap-like structure over the active site of GST M1-1 provides the optimal condition for binding of the xenobiotic substrates mBBr and CDNB, as demonstated by comparing the Km values of these substrates. The wild-type mu class isozyme has the lowest Km value for both xenobiotic substrates amongst the three classes of GSTs (Table 2). The KmCDNB of the pi class isozyme is the highest, while the KmCDNB of the Deletion Enzyme falls between the pi and alpha class values. The alpha and the pi class KmCDNB values are closer to each other than are the alpha and mu class KmCDNB values. This trend is also apparent in comparing the KmmBBr of the three wild-type enzymes, but the KmmBBr of the mu class Deletion Enzyme is higher than those of the wild-type GSTs. Excision of amino acid residues 35–44 results in shortening the mu loop by ~12 Å, as measured in silico; however, there is still a small loop left in position that could provide shielding from the solvent (Fig. 5A,B). The deletion of the mu loop substantially decreases the enzyme's affinity for both xenobiotic substrates, demonstrating that the mu loop is a determinant of the enzyme's affinity for its substrates. The decrease in the enzyme's affinity for CDNB is not as striking as it is for mBBr. Previously, we have shown that CDNB and mBBr occupy two distinct substrate sites (Hearne and Colman 2005). The CDNB substrate site is situated more externally in the active site cleft of GST M1-1, as compared with the mBBr substrate site, which is in a more hydrophobic area of the active site cleft (Hearne and Colman 2005). The positioning of the substrate sites renders the mBBr substrate site more sensitive to solvent exposure than the CDNB site.

In comparison with the wild-type enzyme Vmax value, the Vmax value of the Deletion Enzyme increases approximately threefold with CDNB as the xenobiotic substrate and approximately fivefold with mBBr as the xenobiotic substrate. The excision of amino acid residues 35–44 augments the distance between Tyr115 and Ser209 from 2.92 Å to 3.74 Å. This larger distance may contribute to the increased rate of product dissociation, reflected in the Vmax value. This result may be due to the inability of the Deletion Enzyme to form hydrogen bonds between these two amino acid residues because of the repositioning of the side chain of Tyr115. It has previously been established that Tyr115 hydrogen bonds with the main chain amide nitrogen of Ser209, as well as the side chain hydroxyl of Ser209, interactions that are known to block the channel to the CDNB substrate site and limit segmental motion of the enzyme during catalysis (Johnson et al. 1993; Codreanu et al. 2002). This loss of hydrogen bonding allows for an enhanced rate of product (GS-DNB) release from the active site (Johnson et al. 1993; Codreanu et al. 2002). As in the CDNB catalytic reaction, the loss of hydrogen bonding increases the rate of the physical step of product dissociation, suggesting that it is the rate determining step in the mBBr catalytic reaction.

Previous work has implicated Arg42, Trp45, Lys49, Asn58, and Leu59 in binding the glycyl portion of GSH in the wild-type enzyme (Wilce and Parker 1994). The glycyl portion of GSH is the closest part of GSH to the mu loop. As shown in Figure 5A, Trp35 of the Deletion Enzyme was substantially displaced. In the wild-type enzyme Trp45 points toward the glutathione site and has the potential to hydrogen-bond to the glycyl carboxylate of GSH. In the Deletion Enzyme, however, Trp35 protrudes into the solvent creating an additional 8.7 Å of separation between the indole ring and the glycyl carboxylate, as compared with the distance between these groups in the wild-type enzyme (2.6 Å). The deletion of Arg42 eliminates one of the two positively charged amino acid residues in the vicinity of the glycyl carboxylate of GSH (Fig. 5B); Lys49, as numbered in the wild-type enzyme, is the other positively charged amino acid residue. Lys39 was only slightly displaced in the Deletion Enzyme (0.6 Å shift away from GSH), leaving this positively charged amino acid residue in position (3.6 Å) to interact electrostatically with, or hydrogen bond with, the glycyl carboxylate of GSH. The Deletion Enzyme's Asn48 (not shown) and Leu49 are also only slightly displaced as compared with the wild-type enzyme (not shown). In the Deletion Enzyme (Fig. 6), Leu49 is somewhat closer to the cysteinyl peptide amide of GSH and to Asn48 than in the wild-type enzyme, whereas Asn48 of the Deletion Enzyme is slightly further from the glycyl amide than in the wild-type enzyme. Wilce and Parker (1994) also predicted hydrogen bonding between Gln71, Ser72, and Asp105 (from the opposite subunit) with the {gamma}-Glu-Cys portion of GSH. Gln71 and Ser72 were predicted to hydrogen bond to the {alpha}-carboxylate of GSH, while the side chain carboxylate of Asp105 was predicted to hydrogen bond to the {alpha}-amino group of GSH. The functional oxygen of Gln71 also has the potential to hydrogen bond to this {alpha}-amino group of GSH. In the Deletion Enzyme, the distances between GSH and Gln71, Ser72, and Asp105 change little relative to the wild-type enzyme. Previous work has shown that GSH is secured in the active site of GST M1-1 by at least 15 hydrogen bonds (Ji et al. 1992) and more than a dozen electrostatic interactions (Armstrong 1997). We propose that collectively these shifts in the amino acid residues lining the glutathione site cause a local change in the hydrogen bonding network and electrostatic interactions culminating in a considerable increase in the Deletion Enzyme's Km value for GSH.


Figure 6
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  		Figure 6. Enlargement of the active site of the wild-type enzyme and the Deletion Enzyme showing the proximity of Leu36, Lys39, and Leu49 of the Deletion Enzyme and Leu46, Lys 49, and Leu59 of the wild-type enzyme to the glycyl moiety of GSH. The wild-type enzyme's amino acid residues and its mu loop region are shown in red, the Deletion Enzyme's amino acid residues and its remaining mu loop region are shown in yellow, and GSH is colored by atom (oxygen, red; nitrogen, blue; sulfur, yellow; carbon, white). Please note that amino acid residue 49 of wild-type enzyme is equivalent to amino acid residue 39 of the Deletion enzyme. (In the wild-type enzyme residues 45 to 217 are equivalent to those numbered 10 less in the Deletion Enzyme. Specifically, Trp45 of the wild-type enzyme is equivalent to Trp35 of the Deletion Enzyme.)

 
The pKa obtained from the pH dependence of kcat/KmCDNB has been interpreted as that of the enzyme-bound GSH (Liu et al. 1992; Wang et al. 1996). Examination of the pKa value of the wild-type enzyme and the Deletion Enzyme obtained from the pH dependence of kcat/KmCDNB shows that the ionization of GSH has been perturbed: In comparison with the wild-type enzyme, the pKa value of the Deletion Enzyme is unexpectedly 0.5 pH units lower. Xiao et al. (1996) previously stated that the activation of GSH is mediated through a first sphere interaction with the hydroxyl of Tyr6, as well as second sphere interactions between the main chain amide of Leu12 and an on-face hydrogen bond between the hydroxyl group of Thr13 and Tyr6. In the wild-type enzyme, the oxygen of the hydroxyl group of Tyr6 is 3.2 Å from the thiol of GSH. In the Deletion Enzyme, this distance is increased to 4.1 Å. The only other hydrogen bond donors in close proximity to the sulfur of GSH are two water molecules in the solvation shell (Xiao et al. 1996). Although the side chain of Leu12 is not in an equivalent position in the two enzymes, the backbone amide is nearly equivalent, as shown by in silico modeling. The change in position of the side chain of Leu12 may contribute to a change of environment of the GSH thiol since that substrate has been shown to normally pack against the side chain of Leu12 (Xiao et al. 1996).

The protein region binding the glycyl moiety is the least restrictive domain of the glutathione-binding site of GSTs (Adang et al. 1990). The glycyl portion of GSH is not essential for substrate recognition, orientation, or binding of GSH as is the glutamyl residue; nor is it essential for the catalytic activity of the enzyme, which is dependent on the cysteinyl residue (Adang et al. 1990). The flexibility of the glycyl domain allows for the modification or replacement of the glycyl portion of GSH with various amino acid residues, such as phenylglycine and 4-aminobutyric acid, without affecting the Km (Adang et al. 1990). However, if the glycyl carboxylate is modified or omitted, as in GSHME, the result is a change in both Km and Vmax. We examined the kinetics of the wild-type enzyme and the Deletion Enzyme with GSHME to determine if only the affinity of the enzyme for the glycyl portion of glutathione had been affected by the excision of amino acid residues 35–44. We observed that GSHME exhibits a twofold lower Km value (compared with GSH) for both the Deletion Enzyme and the wild-type enzyme. Since the effect on the KmGSHME is the same for both the wild-type enzyme and the Deletion Enzyme, the groups responsible for the lowering of the Km for the nucleophilic substrate appear to be the same or equivalent in the wild-type and the Deletion Enzyme. The most probable explanation for our finding is that in the wild-type enzyme one oxygen atom of the glycyl carboxylate of GSH interacts with Lys49, while the other oxygen atom hydrogen-bonds with Trp45 and Arg42. In the Deletion Enzyme, Lys39 could provide a similar interaction with the glycyl carboxylate oxygen as provided by Lys49 in the wild-type enzyme (Fig. 6). However, the other oxygen atom of the glycyl carboxylate would be in a different environment, due to the deletion of Arg42 and the reorientation of Trp45, contributing to the increase in Km for GSH. Figure 6 indicates that, when GSHME is substituted for GSH, the lone carbonyl may interact with Lys49 of wild type and Lys39 of the Deletion Enzyme. The potential for hydrophobic interactions between GSHME and the enzyme exists. Leu36 of the Deletion Enzyme is predicted to be the closest residue to the ethyl group of GSHME. This would present the opportunity for hydrophobic interactions to occur. In the model of the Deletion Enzyme, Leu36 is 7.6 Å from GSH; estimating the ethyl group of GSHME to be ~3 Å in length, this interaction is feasible. Leu49 of the Deletion Enzyme is near (5.5 Å) GSHME. Most likely, the increased hydrophobic interactions are responsible for the decrease in KmGSHME relative to KmGSH.

The wild-type enzyme's Vmax value is not affected when GSHME is used as the nucleophilic substrate in place of GSH. In contrast, the Vmax of the Deletion Enzyme decreases when GSHME is used in place of GSH, probably because of the size of glutathione monoethyl ester in comparison to GSH. It has been noted by Adang et al. (1990) that larger analogs of GSH tend to exhibit lower Vmax values. This effect is more prominently seen in the Deletion Enzyme than in the wild-type enzyme due to the potentially smaller volume of the active site during catalysis.

Previously, very little was known about the role of the mu loop in the enzyme's structure and catalytic function. Based on our findings we conclude that the mu loop of GST M1-1 is an important determinant of the enzyme's affinity for its substrates and its stability at 25°C, but it does not have a large effect on the overall structure of the enzyme. Unlike triosephosphate isomerase, a homodimeric enzyme that contains an 11-amino-acid residue loop that closes over the active site when the substrate binds to the enzyme (Joseph et al. 1990; Pompliano et al. 1990), the mu loop of GST M1-1 is not required for the enzyme to exhibit catalytic activity with the substrates examined in this report.


   Materials and methods
TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
References
 
Materials
GSH, S-hexylglutathione immobilized on cross-linked 4% agarose beads, CDNB, and ampicillin were purchased from Sigma Chemical Company. mBBr was purchased from Molecular Probes. The oligonucleotide for mutagenesis and the primers for DNA sequencing were purchased from Biosynthesis, Inc. The Quikchange XL Site-Directed Mutagenesis Kit was obtained from Stratagene and the QIAprep Spin Miniprep Kit was supplied by QIAGEN. All other reagents were purchased from Fisher and were of reagent grade.

Plasmid and mutagenesis
The cDNA encoding rat (Rattus norvegicus) GST M1-1 as well as the 3'-untranslated region of rat GST M1-1 were inserted into a pBR322 vector via the NdeI and EcoRI restriction sites. This plasmid was a generous gift from Ming F. Tam (Institute of Molecular Biology, Academia Sinica, Nankang, Taipei, Taiwan), and was used with the permission of Dr. M. Rosenberg (Smith Kline Beecham), by whom the expression vector pMG27N (of which this is a derivative) was developed.

The following oligonucleotide and its complimentary sequence were used to delete the nucleotides coding for amino acid residues 35–44 in the cDNA of GST M1-1, according to the Stratagene Quikchange XL Site-Directed Mutagenesis kit instruction manual. The codon for the amino acid residue on each side of the deletion is underlined: 5'-GAGATACGCCATGTGGCTGAATGAGAAG-3'.

DNA extraction and purification was completed using the QIAprep Spin Miniprep Kit. DNA sequencing confirmed codon deletion. Sequencing was performed at the University of Delaware Center for Agricultural Biotechnology using an ABI Prism model 377 DNA sequencer (PE Biosystems). The forward sequencing primer is as follows: 5'-ATGCCTATGATACTGGGATA-3' and the reverse primer is 5'-CATTGGGCCAACTTCGAAAA-3'. The mutated DNA was transformed into competent JM105 E. coli cells for expression (Sambrook et al. 1989).

Protein expression and purification
The wild-type enzyme and the Deletion Enzyme were expressed in JM105 E. coli cells. The cells were grown at 37°C in LB media containing 270 µM ampicillin until A600 nm was 0.4–0.6, at which time the cells were induced with a final concentration of 1 mM IPTG. The proteins were expressed at 25°C for 18 h, after which they were harvested by centrifugation at 10,444 g for 25 min at 4°C. The resulting pellet was immediately frozen and stored at –80°C. The cell pellet from 6 L of culture was thawed in a 25°C water bath and resuspended in 50 mL of 10 mM Tris chloride buffer (pH 7.8). The suspension was kept at 4°C. The cells were ruptured by 6 min of sonication (three 2-min intervals of sonication, separated by 30-sec intervals) at 20 kHz and 475 W with a sonicator from Ultrasonic, Inc.

The suspension was centrifuged at 10,886 g for 25 min at 4°C. The supernatant was decanted and applied to a 0.7 x 20 cm column packed with 10 mL of S-hexylglutathione immobilized on cross-linked 4% agarose beads for purification. All column purification procedures were performed at 4°C. Succinctly, the column was equilibrated with 1 L of 10 mM Tris chloride buffer (pH 7.8), and the enzyme suspension was loaded onto the column. The column was first eluted with 1 L of 10 mM Tris chloride buffer (pH 7.8), followed by 0.25 L of 10 mM Tris chloride buffer (pH 7.8) containing 0.2 M NaCl to elute the nonspecifically bound proteins from the column. Both enzymes were eluted with 0.2 L of 10 mM Tris chloride buffer (pH 7.8) containing 2.5 mM S-hexylglutathione, as detected by monitoring the eluate using the standard assay (defined below). The enzyme was dialyzed and concentrated in 0.1 M potassium phosphate buffer (pH 6.5) containing 1 mM EDTA by use of Amicon Ultra Centrifugal Filter Devices (Millipore Corp.), which were spun at 2611g for 15 min at 4°C. The resulting enzyme was rapidly frozen and stored at –80°C. The enzyme concentration was determined with a Hewlett Packard 8453 UV-VIS spectrophotometer, using the extinction coefficient at 270 nm ({Delta}{varepsilon} = 37,700 M–1 cm–1).

Enzymatic assays
The following conditions were used in determining the specific activity (expressed as µmol substrate converted per minute per milligram enzyme) of the enzymes for each set of substrates. The conjugation of 1 mM 1-chloro-2,4-dinitrobenzene (CDNB) and 2.5 mM glutathione (GSH) in 0.1 M potassium phosphate buffer (pH 6.5) containing 1 mM EDTA and 2.5% ethanol was monitored at 340 nm ({Delta}{varepsilon} = 9.6 mM–1 cm–1) at which the increase in absorbance is due to the formation of the product, GS–DNB. This reaction is the standard assay and was monitored for 30 sec using a Hewlett Packard 8453 UV-VIS spectrophotometer (Habig et al. 1974). To evaluate the stability of the enzyme throughout the time of the assay, the reaction was monitored spectrophotometrically for one minute, yielding the same rate as determined for the reaction over 30 sec. This set of conditions was also followed to monitor the reaction of CDNB and glutathione monoethyl ester (GSHME). The reaction of 30 µM monobromobimane (mBBr) and 600 µM GSH in 0.1 M potassium phosphate buffer (pH 6.5) containing 1 mM EDTA and 20% DMF was monitored from the time-dependent increase in fluorescence using a Perkin-Elmer MPF-3 fluorescence spectrophotometer (emission at 480 nm, excitation at 395 nm) (Hulbert and Yakubu 1983). The reaction of mBBr with GSH was monitored over 30 sec, yielding a linear time-dependent change in fluorescence. In each case, the specific activity was corrected for the rate of the spontaneous nonenzymatic conjugation reaction between the xenobiotic substrate and GSH. For all rate determinations the enzyme stock was maintained at 4°C while the assay temperature was 25°C; reaction was initiated by the addition of 20 µL of suitably diluted enzyme solution, followed by 20 µL of GSH to 960 µL of assay solution. The enzyme stock must be kept at 4°C to maintain its stability during the period of the experiment.

To determine the KmCDNB of the wild-type enzyme, a range of CDNB concentrations was used (5 µM–1000 µM), while the GSH concentration was fixed at 2.5 mM. For the Deletion Enzyme, the CDNB concentration range was extended to 2.5 mM and the GSH concentration was held constant at 30 mM. Determination of the KmGSH for the wild-type enzyme in the CDNB–GSH conjugation reaction was accomplished using a range of GSH concentrations (10 µM–2500 µM), while the 1 mM CDNB concentration was constant. For the Deletion Enzyme the CDNB concentration was maintained at 2.5 mM and the range of GSH concentrations was extended to 30 mM. The Km of the wild-type enzyme and the Deletion Enzyme for GSHME was evaluated using a fixed concentration of 2.5 mM CDNB and a range of GSHME concentrations (wild-type enzyme, 20 µM–1000 µM; Deletion Enzyme, 500 µM–5000 µM).

To determine the wild-type enzyme's KmmBBr, a range of mBBr concentrations was used (0.25 µM–60 µM), while the GSH concentration was constant at 600 µM. For the Deletion Enzyme, a range of mBBr concentrations was used (50 µM–500 µM), while the GSH concentration was constant at 600 µM.

The data, in each case of the Km determinations, were fitted to the Michaelis-Menten rectangular hyperbola using SigmaPlot. The Vmax and standard error were calculated from an extrapolation of the data. For all kinetic parameter determinations, the enzyme stock solutions were kept at 4°C, the temperature of the assay was maintained at 25°C and the conditions were generally saturating for the invariant substrate.

The pH dependence of kcat/KmCDNB for the wild-type enzyme and the Deletion Enzyme was determined at 25°C using the following buffers at the indicated pH: 0.1 M sodium acetate at pH 5.0 through pH 5.75 and 0.1 M potassium phosphate containing 1 mM EDTA in the pH range 6.0–8.0. The concentration of CDNB was varied over the range of 10–2500 µM for both enzymes. The reactions were carried out under saturating GSH concentrations for the wild-type GST M1-1 enzyme throughout the pH range 5.75–8.0. In contrast to the wild-type enzyme, evaluation at 10 times the KmGSH of the Deletion Enzyme could not be realized because of the high rate of the nonenzymatic reaction of the substrates. However, the concentration of GSH was at least twofold greater than the KmGSH through the pH range 5.0–7.0 for the Deletion Enzyme. The pH dependence of kcat/KmCDNB was determined by fitting the data to the equation pKa = (kcat/Km)/(1 + 10(pKa–pH)), which was reported by Winayanuwattikun and Ketterman (2005).

The stability of both enzymes was tested in the pH range used to determine the pKa by incubating each enzyme in the buffer used at the indicated pH values for 60 sec. Each enzyme was assayed under standard assay conditions immediately after its addition to the indicated buffer and this rate was recorded. Each enzyme was then assayed in the same manner after 60 sec of incubation in the indicated buffer and this rate was compared with its initial rate.

N-terminal sequencing
The purity of the wild-type enzyme and the Deletion Enzyme was assessed by Edman degradation executed by an Applied Biosystems Procise Sequencing System. The deletion of the 10 amino acid residues was confirmed by 47 reaction cycles of N-terminal sequencing.

Molecular modeling
The amino acid sequence of the Deletion Enzyme was submitted to SWISS-MODEL for homology modeling (Schwede et al. 2005). The model of the Deletion Enzyme was based on the crystal structures of 2GST(B) (Ji et al. 1993), 3GST(B) (Ji et al. 1993), 4GST(B) (Ji et al. 1994), 5GST(B) (Ji et al. 1994), and 6GST(B) (Xiao et al. 1996); (B) indicates that the B subunit was used in modeling. The model was returned as a monomer.

Molecular mass determination
To determine the molecular mass of the monomeric form of each enzyme, the enzymes were dialyzed into water to remove the salt in preparation for electrospray ionization mass spectrometry (ESI-MS). The wild-type enzyme (1.4 x 10–10 M) and the Deletion Enzyme (2.3 x 10–11 M) were each mixed with methanol and 0.1% TFA to assist in aerosol formation and ionization. Each sample was loaded into an AutospecQ (Micromass). Data analysis was performed using the Masslynx software.

Sedimentation equilibrium experiments, run in a Beckman Coulter XL-I analytical ultracentrifuge, were used to determine the weight average molecular weight of the wild type enzyme and the Deletion Enzyme under mild conditions at 0.06 mg/mL and 0.3 mg/mL. Sedimentation equilibrium experiments were performed at speeds of 15,000 rpm, 17,000 rpm, and 20,000 rpm running at 4°C, using an An-50 Ti rotor. After equilibrium was reached, stepwise radial scans at 235 nm for 0.06 mg/mL enzyme samples and 270 nm for 0.3 mg/mL enzyme samples were performed using a step size of 0.001 cm. Enzyme samples in the absence or presence of GSH (wild type enzyme with 2.5 mM GSH and the Deletion enzyme with 5 mM GSH; Sigma) were in 0.1 M potassium phosphate buffer (pH 6.5) containing 1 mM EDTA. The resulting data were fit using the software package IgorPro (Wavemetrics, Inc.) as previously described (Schneider et al. 1997; Kretsinger and Schneider 2003).

Circular dichroism spectroscopy
Circular dichroism spectroscopy was performed on a Jasco J-710 spectropolarimeter, as previously described (Vargo and Colman 2004). Concisely, the ellipticity of the enzyme sample (0.15 mg/mL in 0.1 M potassium phosphate buffer at pH 6.5 containing 1 mM EDTA at 4°C) was measured as a function of wavelength between 200 nm and 250 nm at 0.2 nm increments. The average of five measurements was recorded as the spectrum. Each sample spectrum was corrected for the contribution from 0.1 M potassium phosphate buffer (pH 6.5) containing 1 mM EDTA. For some experiments the CD spectra were recorded at 4°C, 10°C, and 25°C.

Fluorescence spectrophotometry
The steady-state fluorescence spectra of the wild-type enzyme and the Deletion Enzyme (2.2 x 10–5 M enzyme subunits) in 0.1 M potassium phosphate buffer (pH 6.5) containing 1 mM EDTA at 4°C were measured on a Perkin Elmer MPF-3 fluorescence spectrophotometer. The samples were excited at 280 nm and the emission spectrum of each sample was scanned and recorded in the range of 300 nm to 420 nm. The bandwidth for excitation and emission was 10 nm. The spectra were corrected for the background contributed by the buffer. The fluorescence of the enzyme samples was compared with the fluorescence of free tryptophan (8.8 x 10–5 M), free tyrosine (3.1 x 10–4 M), and free phenylalanine (2.6 x10–4 M) at the same molar concentrations at which they are present in the wild-type enzyme sample. There are four tryptophan, 14 tyrosine, and 12 phenylalanine residues in a GST M1 subunit. The concentration of tyrosine in the Deletion Enzyme is 2.9 x 10–4 M since there is one less tyrosine residue in the Deletion Enzyme than in the wild-type enzyme.

Thermostability studies
The thermostability of the wild-type enzyme and the Deletion Enzyme was evaluated by incubating the enzymes (0.3 mg/mL enzyme in 0.1 M potassium phosphate buffer at pH 6.5 containing 1 mM EDTA and 2.5 mM GSH) in a water bath at 4°C, 10°C, or 25°C. An aliquot of the enzyme was removed in a time-dependent manner and assayed at 25°C in 0.1 M potassium phosphate buffer (pH 6.5) containing 1 mM EDTA, 1 mM CDNB, and 2.5 mM GSH. The duration of incubation was 1 h. The activity of the enzymes is expressed as Et/Eo (observed activity at a given time/initial activity). The wild-type GST M1-1 enzyme and the Deletion Enzyme (0.06 and 0.3 mg/mL) in the presence and absence of GSH were also subjected to 76 h of incubation at 4°C to test whether they were stable for the duration of the sedimentation equilibrium experiments.

Synthesis of glutathione monoethyl ester
The procedure used to synthesize GSHME (glycyl carboxylate esterified) was a modification of that of Campbell and Griffith (Campbell and Griffith 1989), which in turn was based on the method of Puri and Meister (1983) with the following exceptions: (1)The reaction vessel was maintained under nitrogen to minimize the oxidation of the reactant (GSH) and the product (GSHME). (2) The reaction was monitored by HPLC (absorbance at 220 nm, 254 nm, and 280 nm); the sample was diluted in solvent A and applied to a Hewlett Packard 1100 RP-HPLC (5 µm, 4.6 mm ID, 250 mm L, Vydac C18) employing a linear gradient from 0% to 30% solvent B over 30 min (where solvent A is 0.1% trifluoroacetic acid [TFA] in water, and solvent B is 90% acetonitrile, 10% water, and 0.1% TFA). The HPLC profiles match those reported by Campbell and Griffith (1989) (data not shown). From their profile we identified peak 2 of our HPLC profile (elution at 11% B) as the glycyl monoethyl ester derivative of GSH. It should be noted that Campbell and Griffith (1989) derivatized the components of the reaction mixture with mBBr (Newton et al. 1981) and monitored the HPLC elutions by fluorescence. We did not derivatize the reaction components; therefore, the elution times in the present study differ from those of Campbell and Griffith (1989). Puri and Meister (1983) identified this product peak as the glycyl monoethyl ester by NMR. (3) The reaction was allowed to proceed for an additional 18 h at 20°C for maximum conversion to product, as recommended by Campbell and Griffith (1989). (4) After precipitation, the product was collected and lyophilized immediately. The glutathione diethyl ester does not precipitate under these conditions. Proper washing of the product is required to remove the diethyl ester. (5) The dry sample was solubilized (20 mg/mL) in solvent A and applied to a RP-HPLC equipped with a Waters 2487 dual absorbance detector (220 nm, 254 nm), Waters 600 pump, and a Linseis L250E recorder (10 µm, 22 mm ID, 250 mm L, Vydac C18) employing a linear gradient from 0% to 100% solvent B (data not shown). This step is essential for purification because traces of the {gamma}-glutamyl monoethyl ester derivative of GSH and the glycyl diethyl ester derivative of GSH are present in the precipitated product. The GSHME peak was collected over liquid nitrogen and shell frozen for lyophilization. A sample of the product was solubilized in solvent A and applied to the Hewlett Packard 1100 RP-HPLC from which the single peak was collected for ESI-MS [(M + H)+ = 336.2].


   Footnotes
 
Reprint requests to: Roberta F. Colman, Department of Chemistry and Biochemistry, University of Delaware, Newark, DE 19716, USA; e-mail: rfcolman{at}chem.udel.edu; fax: (302) 831-6335.

Article published online ahead of print. Article and publication date are at http://www.proteinscience.org/cgi/doi/10.1110/ps.062129506.

Abbreviations: GST, glutathione S-transferase; WT, wild-type GST M1-1; Deletion Enzyme, mu loop (amino acid residues 35–44) deletion mutant GST M1-1 enzyme; GST A1-1, alpha class GST; GST P1-1, pi class GST; GSH, glutathione; GSHME, glutathione monoethyl ester; CDNB, 1-chloro-2, 4-dinitrobenzene; mBBr, monobromobimane, 4-bromomethyl-3,6,7-trimethyl-1,5-diazabicyclo[3.3.0]octa-3,6-diene-2,8-dione; Tris, tris(hydroxymethyl)aminomethane; LB, Luria-Bertani; EDTA, disodium ethylenediamine tetraacetate; DMF, N, N'-dimethylformamide; IPTG, isopropyl-beta-D-thiogalactoside; TFA, trifluoroacetic acid; CD, circular dichroism; ESI-MS, electrospray ionization mass spectrometry; NMR, nuclear magnetic resonance; rpm, revolutions per minute.


   Acknowledgments
 
This work was funded by NIH R01-CA66561 (R.F.C). The Beckman Optima XL-I analytical ultracentrifuge used in this study was obtained and supported by NIH 2P20 RR016472. We thank Dr. Ming Tam and Dr. M. Rosenberg for the plasmid containing GST M1-1, Dr. Melissa A. Vargo for the purified GST A1-1 enzyme, and Luis A. Ralat for the purified GST P1-1 enzyme. We also thank Dr. Joel Schneider for the use of the analytical ultracentrifuge data analysis software, Dr. Yu-Chu Huang for obtaining the N-terminal sequences, Haixia Wu for the ESI-MS data collection and analysis, as well as Lisa A. Haines for her assistance in the synthesis of GSHME.


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 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 References
 
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