The functional implications of the dimerization of the catalytic subunits of the mammalian brain platelet-activating factor acetylhydrolase (Ib)

T.W.P. McMullen1, J. Li1, P.J. Sheffield1, J. Aoki2, T.W. Martin1, H. Arai2, K. Inoue2 and Z.S. Derewenda1,3

1 Department of Molecular Physiology and Biological Physics, University of Virginia, Health Sciences Center, Charlottesville, VA 22906-0011, USA and 2 Department of Health Chemistry, Faculty of Pharmaceutical Sciences, University of Tokyo, Japan


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
The mammalian brain contains significant amounts of the cytosolic isoform Ib of the platelet-activating factor acetylhydrolase (PAF-AH), a unique type of PLA2. This oligomeric protein complex contains three types of subunits: two homologous (63% identity) 26 kDa catalytic subunits ({alpha}1 and {alpha}2) which harbor all the PAF-AH activity, and the 45 kDa ß-subunit (LIS1), a product of the causal gene for Miller–Dieker lissencephaly. During fetal development, the preferentially expressed {alpha}1-subunit forms a homodimer, which binds to a homodimer of LIS1, whereas in adult organisms {alpha}1/{alpha}2 and {alpha}2/{alpha}2 dimers, also bound to dimeric LIS1, are the prevailing species. The consequences of this `switching' are not understood, but appear to be of physiological significance. The {alpha}1- and {alpha}2-subunits readily associate with very high affinity to form homodimers. The nature of the interface has been elucidated by the 1.7 Å resolution crystal structure of the {alpha}1/{alpha}1 homodimer (Ho et al., 1997Go). Here, we examined the functional consequences of the dimerization in both types of {alpha}-subunits. We obtained monomeric protein in the presence of high concentrations (>50 mM) of Ca2+ ions, and we show that it is catalytically inactive and less stable than the wild type. We further show that Arg29 and Arg22 in one monomer contribute to the catalytic competence of the active site across the dimer interface, and complement the catalytic triad of Ser47, Asp192 and His195, in the second monomer. These results indicate that the brain PAF-acetylhydrolase is a unique PLA2 in which dimerization is essential for both stability and catalytic activity.

Keywords: dimerization/mammalian brain/platelet-activating factor acetylhydrolase (Ib)


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Phospholipases A2 (PLA2s) are ubiquitous and diverse enzymes involved in a phethora of physiological phenomena, ranging from phospholipid digestion and venom toxicity to the production and hydrolysis of second messengers and scavenging of biologically active products of oxidative phospholipid degradation. PLA2s are grouped into one superfamily based on the common reaction that they catalyze: the hydrolytic removal of the acyl chain from the sn2 position in phospholipids. The classification within the superfamily, somewhat artificial, relies on the enzymes' requirements for Ca2+, as well as their molecular weights (Dennis, 1994Go, 1997Go). Exhaustive structural information is available with respect to the many members of the low molecular weight, secretory PLA2s (Scott and Siegler, 1994Go). This is a homologous family of Ca2+-dependent enzymes with conserved catalytic mechanism involving a His–Asp dyad, and exhibiting a preference for aggregated substrate (i.e. interfacial activation) (Scott et al., 1990Go). The crystal structure of the cytosolic 85 kDa PLA2 was recently reported at 2.5 Å resolution (Dessen et al., 1999Go). Both the tertiary fold of this enzyme and its catalytic mechanism, which employs an unusual Ser–Asp dyad, are unique among proteins studied to date. Finally, the Escherichia coli outer membrane phospholipidase A (OMPLA) was also characterized structurally by X-ray diffraction, and shown to be a ß-barrel structure with an enzymatic site containing a modification of the trypsin-like catalytic triad with serine, histidine and asparagine as the active residues (Snijder et al., 1999Go).

All of the above PLA2s hydrolyze phospholipids with long acyl chains in the sn2 position. In contrast, platelet-activating factor acetylhydrolases (PAF-AHs) are distinct PLA2s with high substrate specificity towards 1-O-alkyl-2-acetyl-sn-glycero-3-phosphocholine (platelet-activating factor, PAF), where an acetyl group occupies the sn2 position. The physiological function of PAF-AHs is to inactivate PAF, a potent phospholipid messenger molecule with both paracrine and autocrine functions, through a hydrolytic deacetylation of the sn-2 postion, with concomitant formation of the biologically inert lyso-PAF molecule. There are several isoforms, intra- and extracellular (for recent reviews, see Stafforini et al., 1997; Derewenda and Ho, 1999). The plasma (Tjoelker et al., 1995Go) isoform and the so-called isoform II (Hattori et al., 1996Go) are members of the ubiquitous {alpha} hydrolase superfamily (Heikinheimo et al., 1999Go), as inferred from the crystal structure of a bacterial homologue (Wei et al., 1998Go). Consequently, these enzymes are related to neither secretory nor cytosolic PLA2s, but show more similarity to neutral lipases (Derewenda, 1994Go). The third well-characterized mammalian isoform of PAF-AHs is the brain intracellular isoform Ib (Hattori et al., 1993Go). It is an oligomeric complex, which consists of three types of subunits: two homologous {alpha}-subunits, {alpha}1 and {alpha} (63% amino acid identity), which harbor all of the catalytic activity and which are unrelated to any other PLA2s (Hattori et al., 1994aGo, 1995Go), and the ß-subunit, a member of the WD40 family of proteins and the product of the causal gene for Miller–Dieker lissencephaly (Reiner et al., 1993Go; Hattori et al., 1994bGo). The two catalytic subunits associate with high affinity to form a dimer. Interestingly, the composition of this dimer varies during the early development of the organism because of different expression patterns of the {alpha}1 and {alpha}2 proteins: the {alpha}1 is expressed almost exclusively during the fetal stage, whereas {alpha}2 is expressed in adult life (Manya et al., 1998). All three dimers, i.e. the two possible homodimers and the heterodimeric species, are catalytically active, although active-site labeling in the heterodimer suggests that only the {alpha}1-subunit is active in that form. The head-group specificity differs depending on the composition: 1-O-alkyl-2-acetyl-sn-glycero-3-phosphoric acid (AAGPA) is hydrolyzed preferentially by both the {alpha}1-homodimer and the heterodimer, whereas PAF and 1-O-alkyl-2-acetyl-sn-glycero-3-phosphatidylethanolamine (AAGPE) are the preferred substrates for the {alpha}2-homodimer (Manya et al., 1999Go). This pattern of substrate preference is consistent with the notion of the {alpha}1-subunit being the catalytically active one in a functionally asymmetric heterodimer.

In an effort to understand better the structure–function relationships in the intracellular mammalian brain PAF-AH(Ib), we have recently determined the crystal structure of the {alpha}1-homodimer (Ho et al., 1997Go). We reported that the tertiary fold of this protein is unique among PLA2s, with unexpected similarities to the low molecular weight GTPases. The active site contains a classical trypsin-like triad of Ser–His–Asp. More recently, we showed that three residues are largely responsible for the observed strict specificity towards an acetyl moiety, i.e. Leu48, Leu194 and Thr103 (Ho et al., 1999Go). The dimer-forming interface is extensive (1150 Å2 per monomer), and involves 18 key residues, which form a contiguous region around the active site. When the two monomers form a dimer, a donut-shaped structure is created with a deep gorge providing access to two active sites, which are brought to relatively close proximity (Ho et al., 1997Go).

Dimerization is an unusual feature in PLA2s. Although some secretory phospholipases A2 are known to form dimers, the functional implications are not clear. Of all the PLA2s characterized to date, only the E.coli OMPLA requires dimerization for activity, because in a monomer neither the oxyanion hole—required to stabilize the transition states—nor the acyl-binding pocket are formed within a single site. Dimerization places these features of one monomer in proximity of the Ser–His–Asn triad in the other, and so two half-sites in either molecule contribute to the formation of a catalytically competent dimer (Snijder et al., 1999Go). In contrast, the {alpha}-subunits of PAF-AH(Ib) harbor the complete catalytic machinery within a contiguous site in a single monomer, and the functional consequences of dimerization are not intuitively obvious. In this paper, we describe a study designed to probe this issue. Originally our intention was to disrupt the dimer using site-directed mutagenesis, but this approach resulted in the accumulation of insoluble protein in inclusion bodies, suggesting that the monomers are highly unstable and that dimerization is essential for the stability of the protein. We found, however, that in the presence of high Ca2+ concentration both the {alpha}1- and {alpha}2-homodimers dissociate into monomers, which are significantly less stable—as assessed by differential scanning calorimetry—and catalytically inert. Structural analysis of the dimer interface suggested that some residues in one monomer, notably Arg22, Leu26 and Arg29, come into close contact with the loop containing the active site Asp192 and His195 in the other. Our site-directed mutagenesis data show that the interactions of Arg22 and Arg29 across the interface are critical for full expression of activity. This may have important physiological implications. Although differential expression of the {alpha}-subunits is not fully understood, the present study shows that their dimerization is clearly essential for both stability and activity. The inherent instability of monomers and the absence of any evidence of interdimer {alpha}-subunit exchange suggest that the composition of the dimers is effectively regulated exclusively at the transcriptional level.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Protein expression and purification

The cDNAs for both bovine {alpha}-subunits were subcloned into the gluthathione-S-transferase (GST) expression plasmid, pGEX4T1 (Pharmacia), as EcoRI/SalI fragments to produce pGEX4T1-{alpha}1 and pGEX4T1-{alpha}2. These plasmids were individually transformed into the E.coli strain XL1blue (Stratagene). Expression was carried out in liquid cultures in 1–2 l of Luria broth. Growth was initiated at 37°C until an optical density (600 nm) of 0.4–0.6 was reached, and expression was induced by the addition of 0.5–1.0 mM IPTG; growth continued for a further 16 h at 25°C. The cells were harvested by centrifugation (10 min, 5000 g), resuspended in lysis buffer (50 mM Tris–HCl, pH 8.5, 300 mM NaCl, 5 mM EDTA) and lysed by sonication. The soluble fraction was retained after centrifugation (30 min, 15 000 g). GST-fusion proteins were isolated by glutathione Sepharose affinity chromatography. Glutathione Sepharose affinity columns were pre-equilibrated with 20 column volumes of buffer I (50 mM Tris–HCl, pH 8.0, 300 mM NaCl) at room temperature. Soluble protein fractions were allowed to bind to the column for 4–8 h at room temperature with gentle agitation. Columns then drained and washed with 2 l of buffer I overnight at 4°C. Fusion protein was eluted using 20 ml of buffer II (50 mM Tris–HCl, pH 8.0, 500 mM NaCl, 10 mM reduced glutathione). The {alpha}1- or {alpha}2-homodimers were cleaved from their GST tags by thrombin (Sigma) digestion, dialyzed against lysis buffer to remove excess glutathione and GST was removed by passing the resultant GST/{alpha}-subunit mixture again through a glutathione Sepharose column. Pure {alpha}1- or {alpha}2-homodimers were obtained after a final purification step involving size-exclusion chromatography on an S-75 column (Pharmacia, Uppsala, Sweden).

Mutagenesis

cDNA of wild-type bovine {alpha}1-subunit was subcloned into the GST expression vector pGEX4T1 (Pharmacia) as an EcoRI/SalI fragment. The resulting plasmid, pGEX4T1-{alpha}1, was used as the template for site-directed mutagenesis. Mutations were done using the mega-primer polymerase chain reaction (PCR) method (Saiki et al., 1988Go, Higuchi and Ochman, 1989Go). The mega-primer PCR method requires two rounds of amplification, the first with the mutagenic primer in combination with a 5' primer for the pGEX4T1 vector and pGEX4T1-{alpha}1 as template DNA. The PCR product produced was then used as the mega-primer in the second reaction in combination with a 3' pGEX4T1 primer. pGEX4T1-{alpha}1 is used again as template DNA in the second reaction. The second reaction produced the full length, mutated PAF-Ah {alpha}1 gene, flanked by EcoRI and SalI restriction sites, which was then subcloned into pGEX4T1. Positive clones were isolated and verified by DNA sequencing. Mutants were expressed as a GST-fusion protein in E.coli strain BL21(DE3). Cultures were grown and {alpha}1-mutant proteins were isolated and purified as described above.

Differential scanning calorimetry (DSC)

Protein solutions were dialyzed into 20 mM sodium phosphate buffer with 120 mM NaCl and 0.5 mM DTT. For experiments containing calcium, the buffer Tris was substituted for sodium phosphate. The protein concentration varied from ~3.5 to 12.0 mg/ml. The protein concentration was determined from a stock solution of protein using quantitative amino acid analysis. Samples and reference solutions were degassed prior to loading into the calorimeter. All of the calorimetric experiments were performed on an MC-2 differential scanning calorimeter (MicroCal, Northampton, MA). The scan rate was 60°C/h unless noted otherwise and the sample volume was 1.23 ml. The calorimeter was interfaced with an IBM PC and the data were analyzed using Origin 4.1 (MicroCal).

Size-exclusion chomatography

Size-exclusion chromatography was performed on a BioCad Sprint system (PerSeptive Biosystems) using a Superdex 75 column (Pharmacia) capable of separating proteins in the range of 5000–60 000 kDa. A standard buffer (50 mM Tris–HCl, pH 8.0, 200 mM NaCl) was used throughout the purification runs or for the determination of molecular weights. The column was standardized using aldolase, bovine serum albumin, chymotrypsin and ribonuclease. The flow rate was constant at 0.65 ml/min and the amount of protein sample applied to the column varied from 0.01 mg to 0.10 mg. The samples and buffers used to equilibrate the column were identical with those prepared for DSC analysis.

Activity assays

The activity of the {alpha}1- and {alpha}2-homodimer or monomer species was determined in 50 mM Tris–HCl at pH 7.4 with 5 mM EDTA and 20 nmol [3H]acetyl-PAF (DuPont–New England Nuclear) in a total volume of 125 µl. The reaction was allowed to proceed for 30 min at 37°C prior to quenching with 1.25 ml of chloroform—methanol (4:1) and an additional 125 µl of doubly distilled water. An aliquot of 300 µl was taken from the upper phase for radioactivity measurements with a ß-counter (Pharmacia Biotech). For samples preincubated with Ca2+, 100 mM Ca2+ was added to the {alpha}1- or {alpha}2-subunit protein solution and incubated on ice for various times prior to addition of [3H]acetyl-PAF.

X-ray diffraction studies

The Arg22Lys mutant protein was expressed and purified as described above, and crystallized according to the procedure established for wild-type enzyme (Ho et al., 1997Go, 1999Go). Data were collected at the X9B synchrotron beamline (NSLS) at the Brookhaven National Laboratory, and processed using DENZO and SCALEPACK (Otwinowski and Minor, 1997Go). Subsequent calculations were carried out using the CCP4 suite of programs (CCP4, 1994), CNS (Brunger et al., 1998Go) and the program O (Jones et al., 1991Go).


    Results and discussion
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 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
In the {alpha}1/{alpha}1-homodimer structure (Ho et al., 1997Go), the interface is formed primarily by 18 residues of predominantly polar and charged character, including Gln18, Asp20, Arg22, Ser25, Arg29, His106, Arg141, Gln143, Tyr193 and Tyr191. Two salt bridges, Asp20A–Arg141B (A and B denote the two monomers in a functional dimer) and its symmetry-related counterpart, contribute to the stability of the interface. We assumed that the disruption of these specific salt bridges would destabilize the interface sufficiently to yield a monomeric protein. In order to examine the functional significance of the dimerization of the {alpha}-subunits in bovine PAF-AH(Ib), we first designed three site-specific mutants of the {alpha}1-subunit (D20R, D20N and R22E) with the intention of creating a stable monomeric species. However, none of the mutants could be isolated, because of either lack of protein expression or the formation of inclusion bodies. This suggested the possibility that the monomeric protein is unstable, or that the mutants fail to fold properly.

We then used differential scanning calorimetry to probe the stability of the wild-type {alpha}1- and {alpha}2-homodimers, searching for conditions that might induce their dissociation and/or reversible unfolding. The DSC scans for the {alpha}1- and {alpha}2-homodimers as a function of pH are shown in Figure 1Go. The thermally induced unfolding of the {alpha}1- or {alpha}2-subunit from pH 6.5 to 9.5 is irreversible under all conditions examined and the protein precipitated out of solution following denaturation. Below pH 6.0, both {alpha}1- and {alpha}2-homodimers denatured spontaneously and therefore all experiments were conducted above this value. The irreversibility of the thermal denaturation of the {alpha}1- and {alpha}2-subunits of platelet-activating factor acetylhydrolase precluded a formal and detailed analysis of the thermodynamics of unfolding. We therefore limited our calorimetric analysis to the transition temperature and enthalpy of thermal denaturation. For the {alpha}1-homodimer, we found that both the enthalpy and the temperature of unfolding depend, as expected, on pH (Figure 1AGo). From a maximum of 52.3°C at pH 6.5, the temperature of unfolding decreases to 47.5°C as the pH increases to 9.5. The overall enthalpy of the unfolding transition of the {alpha}1-subunit homodimer also decreases as a function of increasing pH, with a maximum value of 164 kcal/mol at pH 6.5 and a minimum value of ~130 kcal/mol at pH 9.5. Similarly, the transition temperature and enthalpy of the thermally induced unfolding of the {alpha}2-subunit homodimers, shown in Figure 1BGo, is also pH dependent. The trend is similar to that shown for the {alpha}1-subunit with both the temperature and enthalpy decreasing as a function of pH. The transition temperature and enthalpy, 51.7°C and 156 kcal/mol, respectively, decrease markedly to 46.2°C and 113 kcal/mol as the pH is increased from 6.5 to 9.5. These parameters did not change in any significant way when the protein concentration was decreased 5-fold, when the scan rate was decreased by a factor of 2–3 or when the ionic strength of the solution was varied (Table IGo). Similar results were obtained with the {alpha}2-homodimers at various scan rates and protein concentrations (results not shown). Overall, the DSC data indicate that the {alpha}1- and {alpha}2-homodimers undergo thermal denaturation and we do not see a stable monomeric species. This was confirmed by size-exclusion chromatography as described later.



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Fig. 1. Comparative DSC scans of the thermal denaturation of the (A) {alpha}1- and (B) {alpha}2-homodimers as a function of pH. Protein solutions contained between 8.5 and 11.0 mg/ml ({alpha}1) and 6.0 and 9.0 mg/ml ({alpha}2) protein in 20 mM sodium phosphate buffer, 120 mM sodium chloride and 0.5 mM DDT. Samples were scanned at 60°C/h and the thermograms were corrected for scan rate and protein concentration. For the thermal denaturation of the {alpha}1-subunits, the {Delta}HvH/{Delta}Hcal ratio is between 0.92 and 1.05 for the samples shown and the thermal unfolding of the {alpha}1-subunits approximates an irreversible, two-state transition ({Delta}HvH = 6.9Tm2/{Delta}T1/2).

 

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Table I. Dependence of wild-type {alpha}1-subunit temperature and enthalpy of unfolding on scan rate, protein concentration and solution ionic strength(Tm ±0.35, {Delta}H ±10%)
 
We then discovered that calcium ions impact on the thermal unfolding of the {alpha}1- and {alpha}2-homodimers. At low calcium concentrations the transition temperature and enthalpy of the unfolding transition of the {alpha}1- and {alpha}2-subunits decrease only slightly as shown in Figure 2Go. However, as the concentration increases above 50 mM, we monitored much larger changes in the temperature and enthalpy of thermal denaturation. For the {alpha}1-subunit, the transition temperature decreases by ~8°C at calcium levels exceeding 250 mM and the enthalpy of the unfolding transition decreases by ~45%. The {alpha}2-subunit demonstrated qualitatively similar calcium-dependent shifts in the transition temperature and enthalpy, but at lower levels of calcium (~100 mM). At all levels of calcium, the thermal denaturation of the {alpha}1- and {alpha}2-subunits of PAF-AH was irreversible. Overall, the temperatures and enthalpies of thermal denaturation of the {alpha}1- and {alpha}2-subunits of PAF-AH in the presence of calcium reveal a qualitatively similar thermodynamic behavior of the subunits. However, the {alpha}2-subunit appears to be more sensitive to calcium given the larger temperature and enthalpy shifts. Taken together, the DSC data suggested the possibility that the homodimers were dissociating in the presence of high Ca2+ concentration into less stable monomers with a melting temperature of ~40°C. To test this hypothesis, we resorted to size-exclusion chromatography, as a means of monitoring the apparent molecular weights in samples treated as described above. Figure 3Go shows the chromatograms of the {alpha}1- and {alpha}2-subunit homodimers, in 150 mM NaCl at pH 7.0, against a selection of molecular weight standards. In both cases, the {alpha}1- and {alpha}2-homodimers exhibit apparent molecular weights of ~60 kDa, consistent with the presence of a dimeric species. This elution profile was not affected either by an increase in pH from 6.5 to 9.5 or by an increase in the salt concentration to as much as 2.5 M (data not shown). The addition of calcium at high concentrations had a major effect on the apparent molecular weights of both the {alpha}1- and {alpha}2-subunits. For the {alpha}1-subunit, the dimer species predominates up to 100 mM Ca2+, but at progressively higher calcium levels the monomer species become prevalent and at 500 mM calcium the {alpha}1-subunit is primarily in monomeric form (Figure 4Go). For the {alpha}2-subunit, the presence of 100 mM calcium is sufficient to shift the dimer–monomer equilbrium decisively in favor of the monomer. Overall, the results of size-exclusion chromatography, consistent with the microcalorimetric experiments described above, indicated that in the presence of 500 mM Ca2+ the {alpha}1- and {alpha}2-subunits are in a monomeric state. This opened up the possibility of examining the monomer for catalytic activity.



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Fig. 2. Comparative DSC scans of the thermal denaturation of the (A) {alpha}1- and (B) {alpha}2-homodimers as a function of increasing calcium concentration. Protein contained between 8.5 and 11.0 mg/ml ({alpha}1) and 6.0 and 9.0 mg/ml ({alpha}2) protein in 20 mM Tris buffer at pH 7.0 with 120 mM sodium chloride and 0.5 mM DDT. Samples were scanned at 60°C/h and the thermograms were corrected for scan rate and protein concentration. For the irreversible thermal denaturation of the {alpha}1-subunits, the {Delta}HvH/{Delta}Hcal ratio falls progressively from near unity to 0.75 at 500 mM Ca2+.

 


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Fig. 3. Size-exclusion chromatography of the {alpha}1- and {alpha}2-homodimers with the molecular weight standards: dextran blue (250 kDa), aldolase (157 kDa) bovine serum albumin dimer (132 kDa), chymotrypsin (25 kDa) and ribonuclease (13 kDa). This series was performed in solution buffered at pH 7.0 by 50 mM Tris with 200 mM sodium chloride. Flow rate was constant at 0.65 ml/min.

 


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Fig. 4. Size-exclusion chromatography of the (A) {alpha}1- and (B) {alpha}2-homodimers as a function of calcium concentration. This series was performed in solution buffered at pH 7.0 with 50 mM Tris and 200 mM sodium chloride. Flow rate was constant at 0.65 ml/min.

 
The above data are consistent with the possibility of a low-affinity Ca-binding site in a protein. However, neither modeling nor attempts to grow crystals of Ca-containing monomers were successful.

The catalytic assays are indicative of the activity of the {alpha}1- and {alpha}2-subunits being highly dependent on the oligomeric state of the enzyme. As the proportion of the enzyme in monomeric state increases at the expense of the dimer with increasing levels of calcium, the relative catalytic activity decreases accordingly (Figure 5Go). In contrast, when the pH is varied from 6.5 to 9.5 the relative activity level of both enzymes remains constant (results not shown). These results are consistent with the notion that amino acids from both monomers contribute to the active site, in such a way that a monomer is catalytically incompetent. Furthermore, the isolated monomer is highly unstable. We therefore set out to identify which specific amino acids from one monomer might contribute to the active site in the other.



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Fig. 5. Enzymatic activity of (A) {alpha}1- and (B) {alpha}2-homodimers as a function of calcium concentration.

 
The crystal structure of the {alpha}1-homodimer (Ho et al., 1997Go) reveals that in each monomer three residues, Arg22, Arg29 and Leu26, come into direct contact with the loop containing two residues of the catalytic triad (Asp192 and His195) in the adjacent molecule. The side chain of Arg22 makes a hydrogen bond to the main chain carbonyl oxygen of the catalytic Asp192, Leu26 is in close van der Waals contact with both C{alpha} and Cß of the same aspartate, and Arg29 creates an H-bonding bridge between the side chain of Tyr193 and the main chain carbonyl of Gly191 (Figure 6a and bGo). All these contacts are likely to stabilize the loop, and consequently the conformations of Asp192 and His195. It should be noted that in acetylhydrolase, in constrast to most other serine hydrolases including trpysin-like proteinases, the catalytic Asp is located on a loop which would be accessible to solvent, and flexible, in a monomer. We therefore hypothesized that if the interactions of this loop across the interface are disturbed, its structural integrity may be seriously impaired. We designed three site-specific mutants to test this hypothesis: these mutants were R22K, R29K and L26A. The R->K mutations were designed to preserve the position charge while removing specific hydrogen bonds, which stabilize the conformation in the triad. The L->A mutation was intended to remove the hydrophobic `cushion' pressed against the side chain of the catalytic aspartate. In each case, we over-expressed the mutant protein and tested the protein for activity.



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Fig. 6. (a) A diagrammatic figure showing the structure of the {alpha}1-homodimer with a highlighted area of the close interactions involving the catalytic loop. (b) A close-up of the detail highlighted in (a), showing the details of the interactions involving Arg22, Leu26 and Arg29, and the catalytic loop in the adjacent monomer (c) The same detail of the structure, in the R22K mutant structure. Note the lack of contact with the catalytic loop, and the different solvent structure.

 
Table IIGo shows the results of the catalytic assays of the mutants. The R22K mutant showed virtually no catalytic activity, whereas the activity of L26A was reduced to ~60% of that of the wild-type. The R29K mutant was only marginally affected. To explore the structural rationale for the observed catalytic impairment of the R22K mutant, we crystallized the protein and determined its three-dimensional structure by X-ray diffraction methods. Fortuitously, the crystals of this mutant diffract to 1.2 Å when placed in a synchrotron X-ray beam, and we were able to collect high-quality data from a single frozen crystal (Table IIIGo). Given that the crystals are isomorphous with those of the wild-type protein, the crystallographic refinement was straightforward, and yielded a highly refined model (conventional crystallographic R-factor 19.2, free R-factor 21.7; see Table IIIGo) which revealed a unique conformation for Lys22, clearly visible in the final 2FobsFcalc electron density map (Figure 7Go). The side chain amide of Lys22 points away from Asp192 in the adjacent monomer, thus breaking the salt bridge interaction observed in the wild-type protein between Arg22 and Asp192. Moreover, the carbonyl of Asp192 is no longer H-bonded to another amino acid, but instead its H-bonding potential is saturated by a new water molecule. Only minor rearrangements occur in the vicinity, the most notable being the distribution of water molecules (Figure 6cGo). Interestingly, the loop carrying Asp192 and His195 is well ordered, with temperature factors not deviating in any significant way from those observed in the wild type structure.


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Table II. Kinetic constants of wild-type and mutant PAF-AH
 

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Table III. Crystallographic data and refinement statistics
 


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Fig. 7. Final 2FobsFcalc electron density map of the R22K mutant, contoured at the level of 1{sigma}. The site of mutation (Arg22->Lys) and adjacent residues and water molecules are shown.

 
The crystal structure of the R22K mutant does not provide any immediately obvious rationale for the dramatic decrease in this mutant's catalytic efficiency. However, several possibilities can be considered. First, it is possible that the side chain of Arg22 provides stabilization to the loop at the stage of the transition state, rather than the ground state. This would imply that in the Michaelis complex the loop ought to show more flexibility. Second, the salt bridge between Arg22 and Asp192 may indirectly affect the pKa of the latter and the function of the catalytic triad. Since the catalytic efficiency of the triad depends on the nature of the Asp–His hydrogen bond, a change in the pKa of the aspartate is likely to have dramatic effects on catalysis. The third possibility, that the arginine is involved in substrate binding rather than catalysis, is unlikely given the position of the side chain of Arg22 in relation to the catalytic Ser47. To assess the likelihood of any, or all, of these mechanisms, further structural studies involving enzyme–inhibitor complexes seem necessary.

Recent literature suggests that the brain isoform of PAF-AH is distantly related to a diverse family of microbial hydrolases, which use complex saccharides and lipids as substrates (Upton and Buckley, 1995Go; Dalrymple et al., 1997Go). While relationships inferred from limited amino acid sequence similarities are sometimes misleading, structural information is more convincing. Indeed, the crystal structure of the rhamnogalacturonan acetylesterase from Aspergillus aculeatus (Mølgaard et al., 2000Go) confirms the notion of evolutionary link between these enzymes. It is not known, however, if dimerization plays any role in the microbial proteins. Among eukaryotes, only Drosophila has been so far shown to harbor a gene coding for a homologue of the {alpha}-subunit of the brain PAF-AH (Sheffield et al., 2000Go). However, the protein is catalytically inert, at least against PAF-related substrates and probably also against other esters, given that two of the three residues in the active site triad are changed. In contrast to the mammalian protein, the Drosophila homologue is largely monomeric, as judged by gel filtration experiments. Hence, the evolutionary origins and roots of substrate specificity, in addition to the phylogeny of gene duplication, continue to be an enigma. As more eukaryotic genome data become available, it will be interesting to see if they contain more clues to the physiological roles of the brain PAF-AH. Our work demonstrates, however, that the dimerization of the mammalian protein is essential for full catalytic activity and stability. In addition, the instability of the {alpha}-monomeric species suggests that the composition of the catalytic {alpha}-dimers is regulated at the transcriptional level and not by dimer rearrangement at the protein level, unless accessory proteins of some kind are involved.

The coordinates of the R29K mutant reported in this paper have been deposited with the Protein Data Bank under accession code 1ES9.


    Notes
 
3 To whom correspondence should be addressed. E-mail: zsd4n{at}virginia.edu Back


    Acknowledgments
 
We gratefully acknowledge Dr Z.Dauter (NCI) for help with data collection and processing. This work was supported by NIH grant NS-36267 to Z.S.D.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Brunger,A.T. et al. (1998) Acta Crystallogr., D54, 905–921.[ISI]

Collaborative Computational Project No. 4, (1994) Acta Crystallogr., D50, 760–763.[ISI]

Dalrymple,B.P., Cybinski,D.H., Layton,I., McSweeney,C.S., Xue,G.P., Swadling,Y.J. and Lowry,J.B. (1997) Microbiology, 143, 2605–2614.[Abstract]

Dennis,E.A. (1994) J. Biol. Chem., 269, 13057–13060.[Free Full Text]

Dennis,E.A. (1997) Trends Biochem., 22, 1–2.[ISI][Medline]

Derewenda,Z.S. and Ho,Y.S. (1999) Biochim. Biophys. Acta, 1441, 229–236.[ISI][Medline]

Derewenda,Z.S. (1994) Adv. Protein Chem., 45, 1–52.[ISI][Medline]

Dessen,A., Tang,J., Schmidt,H., Stahl,M., Clark,J.D., Seehra,J. and Somers,W.S. (1999) Cell, 97, 349–360.[ISI][Medline]

Hattori,K., Adachi,H., Matsuzawa,A., Yamamoto,K., Tsujimoto,M., Aoki,J., Hattori,M., Arai,H. and Inoue,K. (1996) J. Biol. Chem., 271, 33032–33038.[Abstract/Free Full Text]

Hattori,M., Arai,H. and Inoue,K. (1993) J. Biol. Chem., 268, 18748–18753.[Abstract/Free Full Text]

Hattori,M., Adachi,H., Tsujimoto,M., Arai,H. and Inoue,K. (1994a) J. Biol. Chem., 269, 23150–23155.[Abstract/Free Full Text]

Hattori,M., Adachi,H., Aoki,J., Tsujimoto,M., Arai,K. and Inoue,K. (1995) J. Biol. Chem., 270, 31345–31352.[Abstract/Free Full Text]

Hattori,M., Adachi,H., Tsujimoto,M., Arai,K. and Inoue,K. (1994b) Nature, 370, 216–218.[ISI][Medline]

Heikinheimo,P., Goldman,A., Jeffries,C. and Ollis,D.L. (1999) Struct. Fold. Des., 7, R141–R146.[ISI][Medline]

Higuchi,R.G. and Ochman,H. (1989) Nucleic Acids Res., 17, 5865.[ISI][Medline]

Ho,Y.S. et al (1997) Nature, 384, 89–93.

Ho,Y.S., Sheffield,P.J., Masuyama,J., Arai,H., Li,J., Aoki,J., Inoue,K., Derewenda,U. and Derewenda,Z.S. (1999) Protein Eng., 12, 693–700.[Abstract/Free Full Text]

Jones,T.A., Zou,J.Y., Cowan,S.W. and Kjeldgaard,M. (1991) Acta Crystallogr., A47, 110–119.[ISI]

Manya,H., Aoki,J., Watanabe,M., Adachi,T., Asou,H., Inoue,Y., Arai,H. and Inoue,K. J. Biol. Chem., 273, 18567–18572.

Manya,H., Aoki,J., Kato,H., Ishii,J., Hino,S., Arai,H. and Inoue,K. (1999) J. Biol. Chem., 274, 31827–31832.[Abstract/Free Full Text]

Mølgaard,A., Kauppinen,S. and Larsen,S. (2000) Structure, 8, 373–383.[ISI][Medline]

Otwinoski,Z. and Minor,W. (1997) Methods Enzymol., A276, 307–326.[ISI]

Reiner,O., Carrozzon,R., Shen,Y., Wehnert,M., Faustinella,F., Dobyns,W.B., Caskey,C.T. and Ledbetter,D.H. (1993) Nature, 364, 717–721.[ISI][Medline]

Saiki,R.K., Gelfand,D.H., Stoffel,S., Scharf,S.J., Higuchi,R., Horn,G.T., Mullis,K.B. and Erlich,H.A. (1988) Science, 239, 487–491.[ISI][Medline]

Scott,D. and Siegler,P.B. (1994) Adv. Protein Chem., 45, 53–88.[ISI][Medline]

Scott,D.L., White,S.P., Otwinowski,Z., Yuan,W., Gelb,M.H. and Sigler,P.B. (1990) Science, 250, 1541–1546.[ISI][Medline]

Sheffield,P.J., Garrard,S., Caspi,M., Sapir,T., Aoki,J., Inoue,K., Reiner,O. and Derewenda,Z.X. (2000) Proteins: Struct. Funct. Genet., 39, 1–8.[ISI][Medline]

Snijder,H.J., Ubarretxena-Belandia,I., Blaauw,M., Kalk,K.H., Verheij,H.M., Egmond,M.R., Dekker,N. and Dijkstra,B.W. (1999) Nature, 401, 717–721.[ISI][Medline]

Stafforini,D.M., McIntyre,T.M., Zimmerman,G.A. and Prescott,S.M. (1997) J. Biol. Chem., 272, 17895–17898.[Free Full Text]

Tjoelker,L.W. et al. (1995) Nature, 374, 549–552.[ISI][Medline]

Wei,Y., Swenson,L., Castro,C., Minor,W., Derewenda,U., Arai,H., Aoki,J., Inoue,K., Servin-Gonzalez,L. and Derewenda,Z.S. (1998) Structure, 6, 511–519.[ISI][Medline]

Upton,C. and Buckley,J.T. (1995) Trends Biochem. Sci., 20, 178–179.[ISI][Medline]

Received June 1, 2000; revised August 9, 2000; accepted September 15, 2000.