Site-directed mutation of conserved cysteine residues does not inactivate the Streptococcus pyogenes hyaluronan synthase

Coy D. Heldermon, Valarie L. Tlapak-Simmons, Bruce A. Baggenstoss and Paul H. Weigel1

Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK 73190, USA

Received on May 30, 2001; revised on August 17, 2001; accepted on August 29, 2001.


    Abstract
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Hyaluronan synthase (HAS), the enzyme responsible for the production of hyaluronic acid (HA), is a well-conserved membrane-bound protein in both prokaryotes and eukaryotes. This enzyme performs at least six discrete functions in producing a heterodisaccharide polymer of several million molecular weight and extruding it from the cell. Among the conserved motifs and domains within the Class I HAS family are four cysteine residues. Cysteines in many proteins are important in establishing and maintaining tertiary structure or in the coordination of catalytic functions. In the present study we utilized a combination of site-directed mutagenesis, chemical labeling, and kinetic analyses to determine the importance of specific Cys residues for catalysis and structure of the HA synthase from Streptococcus pyogenes (spHAS). The enzyme activity of spHAS was partially inhibited by cysteine-reactive chemical reagents such as N-ethylmaleimide. Quantitation of the number of Cys residues modified by these reagents, using MALDI-TOF mass spectrometry, demonstrated that there are no stable disulfide bonds in spHAS. The six Cys residues of spHAS were then mutated, individually and in various combinations, to serine or alanine. The single Cys-mutants were all kinetically similar to the wild-type enzyme in terms of their Vmax and Km values for HA synthesis. The Cys-null mutant, in which all Cys residues were mutated to alanine, retained ~66% of wild-type activity, demonstrating that despite their high degree of conservation within the HAS family, Cys residues are not absolutely necessary for HA biosynthesis by the spHAS enzyme.

Key words: cysteine residues/disulfide bonds/HA biosynthesis/mutagenesis/synthase


    Introduction
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
In vertebrates hyaluronic acid (HA) is a key component of extracellular matrices, cartilage, and vitreous compartments (Evered and Whelan, 1989Go; Laurent and Fraser, 1992Go; Knudson and Knudson, 1993Go; Toole, 1997Go; Abatangelo and Weigel, 2000Go) and plays key roles in development and fertilization (Fenderson et al., 1993Go; Lin et al., 1994Go). In some bacteria, such as Group A and Group C streptococcus, HA acts as a virulence factor by forming an extracellular capsule that is immunoprotective, enabling these cells to evade host defenses (Whitnack et al., 1981Go; Wessels et al., 1991Go, 1994). The family of enzymes responsible for HA production in viral (DeAngelis et al., 1997Go), bacterial (Kumari and Weigel, 1997Go; DeAngelis et al., 1998Go; Ward et al., 2001Go), and vertebrate organisms (Itano and Kimata, 1996aGo,b; Spicer et al., 1996Go; Shyjan et al., 1996Go; Watanabe and Yamaguchi, 1996Go) was identified after the initial identification and cloning of Streptococcus pyogenes hyaluronan synthase (spHAS), the Group A streptococcal HA synthase (HAS) enzyme (DeAngelis et al., 1993aGo,b). The vertebrate enzymes can be grouped into three evolutionarily conserved subfamilies (Weigel et al., 1997Go), all of which share about 30% identity with the three streptococcal HA synthases, Streptococcus equisimilis HAS (seHAS) (Kumari and Weigel, 1997Go), Streptococcus ubris HAS (suHAS) (Ward et al., 2001Go), and spHAS (DeAngelis et al., 1993bGo). Another bacterial HAS from Pasteurella multocida is very distinct in its properties and primary structure from every other HAS discovered thus far (DeAngelis et al., 1998Go), and was therefore designated a Class II HAS (DeAngelis, 1999Go).

We previously purified the recombinant spHAS and seHAS enzymes, expressed in Escherichia coli (Tlapak-Simmons et al., 1999aGo) and characterized both enzymes with respect to their kinetic constants (Tlapak-Simmons et al., 1999bGo), functional size (Tlapak-Simmons et al., 1998Go), and their requirement of a phospholipid, particularly cardiolipin, for enzyme activity (Tlapak-Simmons et al., 1999aGo). Although the streptococcal HASs are relatively small at <49 kDa, they mediate at least six discrete functions: the ability to bind two different sugar nucleotide precursors, to catalyze two distinct glycosyltransferase reactions, to bind the HA acceptor polymer, and to translocate the growing HA chain through the enzyme and the cell membrane.

We recently found that spHAS and seHAS are sensitive to sulfhydryl reagents, such as iodoacetamide or NEM, which partially inhibit enzyme activity (Kumari et al., unpublished data). This inhibition implicates the involvement of one or more Cys residues in at least one of these six distinct activities of HAS required for HA biosynthesis. There are six Cys residues in spHAS, four of which are conserved perfectly in seHAS and suHAS (Figure 1); both of these latter enzymes have only four Cys residues (Kumari and Weigel, 1997Go; Ward et al., 2001Go). These four Cys residues in turn are generally conserved among the three vertebrate HAS isoenzymes (Weigel et al., 1997Go). No one has yet addressed the possibilities that one or more of these conserved Cys residues within the Class I HAS family may be critical for enzyme activity or may participate in the formation of disulfide bonds.



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Fig. 1. Conservation of Cys residues within the Class I HAS family. The regions shown for these typical HAS family members correspond to the four Cys residues that are identical among the three streptococcal HAS proteins. All Cys residues are in bold. The four conserved streptococcal Cys residues and those that are identical or proximal in the other HAS family members are also shaded in gray. The arrow indicates the spHAS Cys225 residue that is conserved in all family members. Other residues identical in the three streptococcal proteins and in other HAS proteins are highlighted in gray. The eukaryotic sequences shown are from human (hsHAS), mouse (mmHAS), chicken (ggHAS), Xenopus laevis (xlHAS) and chlorella virus (cvHAS).

 
In the present study we used a combination of chemical reagent accessibility, site-directed mutagenesis, inhibition by N-ethylmaleimide (NEM), and mass spectroscopy to demonstrate that there are no stable disulfide bonds in spHAS and that none of the Cys residues in this HAS are necessary for enzymatic activity.


    Results
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Mutagenesis of Cys residues and expression of spHAS
To explore the possible presence of disulfide bonds and the functional roles of the conserved Cys residues in the enzymatic activity of the S. pyogenes HAS, each of the six Cys residues in spHAS was mutated to Ser or Ala. Subsequently, we produced spHAS mutants with combinations of Cys-to-Ala changes, by using site-directed mutagenesis or restriction enzyme digestion and ligation of HAS fragments from different mutants. Studies with crude membranes, in which the enzyme activity of spHAS mutants were initially normalized to total membrane protein, indicated that alteration of some Cys residues had a dramatic affect on HA production. For example, spHAS(C225A) appeared to be nearly inactive, and spHAS(C261A) and spHAS(C280A) had less than half the activity of wild type. However, these initial impressions were incorrect due to significant variations in the expression of spHAS protein among the various mutants.

Therefore, to normalize for the level of HAS protein expression, we developed a sensitive and quantitative western blot–based assay (Heldermon et al., 2001Go). Because all of our HAS constructs contain a C-terminal His6 tag, which is efficiently recognized by a commercial anti-His5 monoclonal antibody, we used this antibody after biotinylation as the primary antibody for analysis of western blots followed by incubation with 125I-streptavidin as the secondary reagent. Unlike standard western analysis, this detection protocol provides greater sensitivity as well as the ability to quantitate HAS protein over a much broader concentration range. The normalizations for HAS protein expression were performed relative to known amounts of purified spHAS-His6 included in each analysis as internal standards. Based on the normalized results, it was clear that spHAS(C225S) was expressed at the lowest level relative to any of the other mutants, ~66% of wild type (Table I). The protein expression levels for the majority of single Cys-mutants were not significantly different than wild type, although the spHAS(C124S) and spHAS(C261A) variants may have been elevated by ~35% (P ~ 0.05). Interestingly, most of the multiple Cys-mutants as well as the Cys-null mutant were expressed at three- to fivefold higher levels than the wild-type enzyme. These differences in relative expression of these spHAS variants were consistent in multiple experiments, with independent cell growth and enzyme induction, indicating that several of the Cys residues in spHAS, particularly the conserved Cys at position 225, may influence the initial folding and stability of the enzyme.


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Table I. Expression levels of various Cys-mutants of spHAS.
 
Enzymatic analysis of spHAS mutants
Kinetic analyses of the single and multiple spHAS Cys-mutants were performed to investigate the possibility that multiple Cys residues are critical in a coordinated way for enzyme activity. The activity of each of the mutants was assayed and normalized by the above method to determine its maximum velocity (Vmax) and Michaelis-Menton (Km) constants for UDP-GlcUA and UDP-GlcNAc (Figures 2 and 3; Table II). This analysis revealed no dependence of HAS activity on any single Cys residue. These assays also revealed no extreme changes in maximal enzyme activity relative to wild-type spHAS. The spHAS(C225S) and spHAS(C280A) mutants had the most reduced activities with Vmax values at 30–50% of wild type. The spHAS(C261,280A) and Cys-null mutant had 50–75% of the wild-type activity. Interestingly, spHAS(C124,366,402A) and spHAS(C366A) had an increased activity that was ~150% of wild type. The other single mutants, as well as spHAS(C124,402A) and spHAS(C124,261,280,366,402A), demonstrated less than a 25% variation from the wild-type Vmax.



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Fig. 2. Kinetic analysis of UDP-GlcUA utilization by Cys-mutants of spHAS. Membranes prepared from cells expressing the indicated spHAS mutant were assayed as described in Materials and methods to assess the Michaelis-Menton constants for UDP-GlcUA: wild type (squares), C124,366,402A (base-up triangles), C124,261,280,366,402A (base-down triangles), and the Cys-null mutant (circles).

 


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Fig. 3. Hill analysis of UDP-GlcNAc utilization by Cys-mutants of spHAS. Hill plots of data obtained from Km assays of wild type and several mutant spHAS proteins, performed as in Figure 2, demonstrate that the cooperative nature of UDP-GlcNAc utilization is not affected by alteration of Cys residues. The spHAS variants shown are: wild type (squares), C124,366,402A (base-up triangles), C124,261,280,366,402A (base-down triangles), and the Cys-null mutant (circles).

 

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Table II. Kinetic constants for Cys-to-Ser/Ala mutants of spHAS
 
All the mutant spHASs were also within 25% of the wild-type enzyme for their KUDP-GlcNAc values. Interestingly, there were no changes in the sigmoidal behavior for UDP-GlcNAc utilization by any of the mutant enzymes. When these data were analyzed using the method of Hill (1913), the Hill numbers were all ~2 (Table II), which indicates a high degree of cooperativity associated with the ability of all the mutant enzymes to bind and use UDP-GlcNAc at a fixed UDP-GlcUA concentration. Thus, the cooperativity observed for the utilization of UDP-GlcNAc by the spHAS enzyme (Tlapak-Simmons et al., 1999bGo) is not influenced by or dependent on any of its six Cys residues. Similarly, none of these Cys residues contribute structurally or otherwise to a possible secondary binding site for UDP-GlcNAc, specifically, an allosteric binding site.

The KUDP-GlcUA values for all of the single Cys-mutants and the two double Cys-to-Ala mutants were within 50% of wild type. The remaining multiple Cys-to-Ala mutants exhibited KUDP-GlcUA values that were two to three times that of wild type. Although these multiple Cys-mutations do alter the activity of the enzyme by decreasing the efficiency of utilizing UDP-GlcUA, they do not do so in a large way. Furthermore, the relatively modest difference in activity between the Cys-null mutant and wild type spHAS clearly shows that cysteine residues are not absolutely necessary for HA synthesis, either catalytically or structurally.

Inhibition of spHAS activity by NEM
Our initial finding that the wild-type streptococcal HAS enzymes were inhibited by NEM (Kumari et al., unpublished data) was the impetus for investigating the role of Cys residues in the enzyme. NEM treatment of membranes from a panel of multiple Cys-mutants showed that this inhibition was no longer present in spHASCys–null or the mutant with only Cys225 intact, whereas NEM sensitivity remained in the other multiple Cys-mutants (Figure 4). These results indicate that the inhibition of the wild-type enzyme by NEM or other sulfhydryl-reactive agents is most likely due to modification of the Cys residues alone, rather than the loss of the S-H group. The lack of inhibition of the single Cys-containing mutant demonstrates that Cys225 is either predominantly inaccessible to modification by NEM due to its position in the enzyme or that this particular cysteine residue is not involved in the inhibition response of the enzyme when modified by NEM.



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Fig. 4. Inhibition of wild-type and Cys-mutants of spHAS by NEM. The activity of the indicated spHAS enzymes in membranes was assessed after pretreatment with (gray bars) or without (black bars) 20 mM NEM at room temperature for 90 min. Wild type and many of the Cys-mutant variants of spHAS with multiple Cys residues mutated are still sensitive to NEM inhibition. The mutant containing only one Cys residue at Cys225 and the Cys-null mutant were not sensitive to NEM inhibition.

 
Assessment of disulfide bond formation in spHAS
Although Cys residues may not be required for the enzymatic activity of the HAS proteins, they could still be important in the structural integrity and long-term stability of the enzyme as indicated by the reduced expression of the spHAS(C225S) mutant and the increased expression of the spHAS(C124S) and spHAS(C261A) mutants (Table I). Cys residues could also be important for maintaining the proper enzyme conformation to allow extrusion of the growing HA chain through the membrane. The primary way in which Cys residues play structural roles in proteins is by forming either inter- or intramolecular disulfide bonds. To investigate the possibility of disulfide bonds in spHAS, we utilized a chemical labeling approach to determine the number of Cys residues that are free and, therefore, could not be involved in disulfide bonding. (+)-Biotinyl-3-maleimido-propionamidyl-3,6-dioxaoctanediamine (biotin-PEO-maleimide) was allowed to react with purified spHAS while bound to a Ni2+-loaded nitrilotriacetic acid (NTA) column, and the modified protein products were then analyzed by matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS) (Figure 5). In nondenaturing conditions, treated wild-type spHAS samples revealed a distribution of derivatized products with increased masses equal to the addition of from one-to-six biotin-PEO-maleimide groups per spHAS, with the majority of the protein being modified by the addition of five or six groups (Figure 5A). The masses observed for all of the adducts differed from the predicted values by <0.03%. As a control for this chemical modification approach, samples of spHASCys–null were treated with biotin-PEO-maleimide in the same way to verify that no derivitized enzyme products would be formed in the absence of Cys groups. The result (Figure 5B) demonstrates that no covalent adducts form with the Cys-null protein, which confirms that the modifying reagent is specific for Cys and does not react with any other amino acid side chains. Treatment of the wild-type or Cys-null spHAS proteins with biotin-PEO-maleimide in the presence of 6 M guanidinium hydrochloride gave essentially the same results as obtained in the absence of the denaturing agent, although the degree of modification was slightly greater (not shown). This latter result indicates that spHAS contains no weak disulfide bonds that might be susceptible to reversible reduction when the protein is denatured. The overall results demonstrate that there are no disulfide bonds in the wild-type spHAS enzyme, and that there is a mixed degree of exposure of the six Cys residues in this protein to the biotin-PEO-maleimide reagent in solution.




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Fig. 5. MALDI-TOF mass spectrographs of spHAS-His6 covalently modified by a sulfhydryl reagent. Wild-type spHAS-His6 (A) or the Cys-null mutant of spHAS-His6 (B) were bound to Ni2+ -NTA resin, washed and incubated for 2 h at 4°C with (upper traces in each panel) or without (lower traces in each panel) 10 mg/ml biotin-PEO-maleimide. The columns were washed and the proteins were then eluted and prepared for mass analysis as described in Materials and methods. The centroid mass-to-charge ratios are indicated above the observed peaks and the predicted mass-to-charge ratios for covalent adducts containing three, five, or six biotin-PEO-maleimide groups per wild-type enzyme molecule are indicated in parentheses. The centroid (and predicted) m/z ratios for the protein adducts containing two or four biotin-PEO-maleimide groups per wild-type enzyme molecule, which were omitted for clarity, were 49,740.0 (49,727.9) and 50,764.9 (50,779.1), respectively. The predicted m/z ratio for the (MH)+ ion of unmodified spHASCys–null-His6 (with six Ala residues replacing the six Cys residues) is 48,484.4.

 

    Discussion
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Cys residues can be important determinants of protein function both structurally and catalytically. Despite the ability of Cys-reactive agents to inhibit HA synthesis by the wild-type streptococcal HASs as described here and in another study (Kumari et al., unpublished data), this is the first report that cysteine residues are not necessary for the catalytic activity of an HAS. Even with all Cys residues changed to Ala, spHAS was still able to carry out the initiation and extension of HA chains with a specific activity and Km values for the two substrates similar to the wild-type enzyme. Furthermore, their susceptibility to chemical derivitization indicated that the six Cys residues present in the wild-type spHAS are not involved in the formation of stable disulfide bonds. In the wild-type spHAS or most of the Cys-mutants described here, the presence of a new group (S-R) after modification by NEM may create steric hindrance for one or more of the functions needed for catalysis. For example, movement of a region of the protein involved in one of the two glycosyltransferase functions might be altered so it becomes less efficient (resulting in a lower Vmax) when a bulky R group is present. In support of this hypothesis, preliminary evidence suggests that a greater inhibitory effect occurs when Cys residues are modified with larger R groups (Kumari et al., unpublished data).

Finally, despite the high degree of conservation of four of the six Cys residues within the larger HAS family and their complete conservation among the three streptococcal HAS enzymes (Figure 1), the spHASCys–null mutant was completely insensitive to inhibition by NEM or other sulfhydryl-reactive reagents. Cys225 is the best candidate Cys residue to explain the NEM sensitivity of spHAS, seHAS, and probably the other HAS enzymes as well, because this Cys residue is the only one that is absolutely conserved in the Class I HAS family (Figure 1). A surprising result, therefore, was that the quintuple-Cys mutant of spHAS containing only Cys225 was not inhibited by NEM or other -SH modifying agents. Although spHAS is functional even with Cys225 replaced by Ser or Ala, ongoing experiments (Kumari et al., unpublished data) indicate that this highly conserved residue may be near or within one of the sugar nucleotide binding sites and may participate in HA biosynthesis in an as-yet-undefined way.

Based on all of the above results we conclude that the role of the Cys residues in the structure or catalysis of the streptococcal HAS proteins must be subtle, because replacement of these residues with Ala does not substantially inhibit the enzyme’s activity or, therefore, presumably its structure. Because the Cys-null mutant is active, this subtle structural role cannot be essential unless there is a substantial difference in behavior of this mutant enzyme in live cells compared to isolated membranes or the detergent-solubilized, purified protein. Studies are in progress to test this possibility.

One possible functional role for the Cys residues in spHAS, which would not be revealed by the present analysis, is in the control of HA product size. It is possible for the HAS enzymes in general that distinct mechanisms control HA chain product length, that is, the size distribution of the HA chains produced, and the overall HA biosynthesis capacity (Abatangelo and Weigel, 2000Go). Another important consideration in evaluating the importance of Cys residues in spHAS (and the other Class I HAS family members in general) may be their involvement in HA translocation. The mechanism by which these enzymes are able to hold onto the growing HA chain while they continuously extrude the polysaccharide through the bacterial cell or plasma membrane is still unknown. We refer to this extrusion process as a translocation, because the HA is not completely transferred across the membrane and released as would occur in a typical transport process. The synthesis and extracellular accumulation by some bacteria of polysaccharides, such as polysialic acid, often requires multiple factors and proteins encoded by very complex multigene operons (Moxon and Kroll, 1990Go; Bliss and Silver, 1996Go). In contrast, all of the genetic and biochemical evidence to date (reviewed in Weigel, 1998Go) demonstrate that the streptococcal enzymes are able to initiate HA chain formation and then rapid extension of the HA chain in the absence of any primer or other proteins. Other than the two sugar nucleotide substrates and Mg2+, the purified spHAS or seHAS enzymes only require a phospholipid (Tlapak-Simmons et al., 1999aGo) to produce high-molecular-weight HA (>106 Da). In particular, cardiolipin dramatically stimulates the specific activity of detergent solubilized or purified spHAS and seHAS. The size distribution of HA products is very similar for enzyme in isolated membranes or after solubilization with dodecylmaltoside and affinity purification (data not shown). Therefore, the presence of a natural intact phospholipid bilayer and membrane does not affect the ability of the HAS enzymes to synthesize HA. Presently we do not have a suitable assay to evaluate the ability of the wild-type or Cys-mutant enzymes to translocate HA.

The creation of a spHASCys–null mutant that retains enzymatic activity should enable a more in-depth analysis of the tertiary structure of spHAS and possible conformation changes during substrate binding, catalysis, or HA translocation. To understand these processes, it is necessary to determine the molecular proximity and interactions of various domains within the protein in a more defined way. For example, as reported for the Lac permease (Frillingos et al., 1998Go), Cys-scanning mutants of spHAS containing a single unique Cys residue at a desired position may enable one to perform electron paramagnetic resonance studies by modifying this Cys residue with a suitable probe to determine the proximity of that residue to another region of the molecule (Voss et al., 1997Go). Similarly, chemical modification of a single unique Cys residue with a fluorescent probe may enable analysis of the localized environment within different regions of the protein (Jung et al., 1994Go). Finally, Cys-scanning or site-specific mutagenesis followed by assessment of possible disulfide bond formation (Wang and Kaback, 1999Go) could help establish interacting or proximal domains within HAS. These approaches, based on the present finding that spHASCys–null is active, hold promise for elucidating the structure and function of spHAS and furthering our understanding of the synthesis of large glycosidic polymers.


    Materials and methods
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Vectors, primers, and reagents
pEx-1 was purchased from Promega as part of the Altered Sites Mutagenesis kit. Successful mutagenesis was achieved with the Quick Change Mutagenesis kit from Stratagene. Primers were synthesized by the Great American Gene Company, NBI, or Midland Certified Reagent Company. Nonradiolabeled UDP-GlcUA, and 2,4,6-trihydroxyacetophenone were from Fluka. UDP-GlcNAc was from Sigma. UDP-[14C]GlcUA was from New England Nuclear. Agarose was from Bio-Rad. Biotin-PEO-maleimide was from Pierce Chemical. NEM and all other reagents were from Sigma unless otherwise noted. To confirm the entire open reading frame (ORF) of HAS mutants, DNA sequencing was performed either using the T7 or polymerase chain reaction sequencing kits from Amersham or by the microsequencing facility operated by the Department of Microbiology and Immunology at the University of Oklahoma Health Sciences Center. Anti-His5 monoclonal antibody and Ni2+-NTA resin were from Qiagen.

Mutagenesis of Cys residues
Single mutants were generated by the Altered Sites Mutagenesis or Quick Change Mutagenesis protocols using primers (Table III) designed to change the Cys residues at positions 124, 225, 261, 280, 366, or 402 of spHAS containing His6 at the C-terminus (Tlapak-Simmons et al., 1999aGo). After generating and confirming the entire sequence of each spHAS(Cys-to-Ser) mutant produced in the Altered Sites vector, internal restriction sites within the HAS ORF were used to transfer mutated regions to the spHAS insert in pKK223 (this vector carrying HAS is designated pKK3K). Cys-to-Ala mutants of spHAS were generated directly in the pKK3K vector using the Quick Change mutagenesis method. Site-directed mutagenesis was used to generate the C124,C402A double mutant, and then C366A was added by restriction fragment exchange to generate a triple mutant. Site-directed mutagenesis was also used to create the double mutant spHAS(C261A,C280A). The mutants containing five or six mutated Cys residues were generated by utilizing restriction sites to combine fragments of spHAS containing different mutations. For example, AvrII and MfeI were used to combine the spHAS(C124A,C366A,C402A) triple Cys-mutant and the spHAS(C261A,C280A) double mutant to create spHAS with only Cys225 intact. Finally, BglII and AvrII were used to splice spHAS(C225A) into the latter quintuple Cys-mutant to generate the Cys-null clone, designated spHASCys–null. All Cys-to-Ala/Ser mutants were confirmed over the full ORF by automated DNA sequencing.


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Table III. Oligonucleotides for Cys-to-Ser/Ala site-directed mutagenesis of spHAS
 
Determination of recombinant spHAS-H6 content in membranes
Membranes were isolated from E. coli SURE strains expressing mutant or wild-type spHAS, fractionated by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) according to the procedure of Laemmli (1970)Go and the proteins were electrotransferred to nitrocellulose as described by Towbin et al. (1979)Go with minor modifications (Tlapak-Simmons et al., 1998Go). Western blot analysis was performed by incubating these blots for 1 h at room temperature with a biotinylated anti-His5 monoclonal antibody, as the primary antibody, then washing and incubating with 3 µg/ml 125I-streptavidin, prepared as described earlier (Heldermon et al., 2001Go). After 1 h at room temperature, the blots were washed, dried, and exposed to a Phosphoscreen for 1–72 h. The screens were analyzed using a Molecular Dynamics Phosphoimager and integrated density values were obtained for each spHAS band. Integrated density values were also obtained for increasing amounts of affinity-purified spHAS (Tlapak-Simmons et al., 1999aGo), which was used as an internal standard in the same blot to generate a standard curve. Dilutions of membrane samples were made as necessary to ensure that all estimates of HAS quantity were in the linear range of the assay. Those membrane samples giving a linear response with increasing protein were then compared to the standard curve to calculate the amount of spHAS present per µl membrane suspension. These values were then used to normalize the kinetic results obtained using membranes from the various spHAS mutants. A typical standard curve for this assay can be found in Figure 2 of Heldermon et al. (2001)Go.

Enzymatic analysis of HAS mutants
E. coli SURE cells, previously transformed with plasmids containing wild-type or mutant spHAS genes, were grown to an A600 of ~1.2 and induced with 1 mM isopropyl thio-ß-D-galactoside for 3 h. Cells were harvested, and membranes were prepared as described previously (Tlapak-Simmons et al., 1999aGo). The activities of mutant spHAS variants were assessed by measuring their Vmax and Km values in isolated membranes, normalized as described above for the amount of enzyme expressed. The Km values were determined using a descending paper chromatography assay (Tlapak-Simmons et al., 1999bGo), holding one UDP-sugar substrate constant and varying the other from 0.01 to 4 mM. Data were analyzed by linear regression using Haynes-Wolf plots for UDP-GlcUA or Hill plots for UDP-GlcNAc.

Inhibition of spHAS activity by NEM
Membrane preparations from wild-type and various spHAS mutants (i.e., C124,402A, C261,280A, C124,366,402A, C124,261,280,366,402A, and the Cys-null mutant) were incubated in 50 mM sodium, potassium phosphate, pH 7.0, 75 mM NaCl and 10% (v/v) glycerol with or without 20 mM NEM for 90 min on ice. The ability of the membrane samples to synthesize HA was then assessed by adding the following to the final concentrations indicated: 1 mM UDP-GlcUA, 1 mM UDP-GlcNAc, 0.68 µM UDP-[14C]GlcUA in 25 mM sodium/potassium phosphate, pH 7, 75 mM NaCl, 10 mM MgCl2, 1 mM dithiothreitol, 0.1 mM ethylene glycol bis(ß-aminoethyl ether)-N,N,N',N'-tetraacetic acid, 15% glycerol. Reactions were shaken for 1 h at 30°C in a Taitec E-36 micromixer, stopped by the addition of SDS to 2% (w/v), and spotted onto No. 3MM Whatman paper for descending paper chromatography overnight using 1 mM ammonium acetate pH 5.5:ethanol (7:13). [14C]GlcUA incorporation into high-molecular-weight HA was assessed by liquid scintillation spectroscopy to determine the radioactivity remaining at the origin. Confirmation that the latter material is authentic HA was obtained by showing its complete loss after treatment with streptomyces hyaluronidase.

Assessment of disulfide bond formation
Wild-type spHAS-His6 was bound to a Ni2+ chelate column (Qiagen) and washed as previously described (Tlapak-Simmons et al., 1999aGo). While still bound to the resin, the enzyme was incubated with biotin-PEO-maleimide (10 mg/ml) in the presence or absence of 6 M guanidinium-HCl for 2 h at 4°C. The column was washed, and spHAS-His6 was eluted with distilled water containing 0.5% (v/v) trifluroacetic acid and 0.02% (w/v) dodecylmaltoside. To assess the degree of modification of Cys residues, samples containing purified spHAS were analyzed by MALDI-TOF MS using a Voyager Elite mass spectrometer (Applied Biosystems, Framingham, MA), which was equipped with a N2 laser (337 nm), located in the NSF EPSCoR Oklahoma Laser Mass Spectrometry Facility. A 1-µl aliquot of sample was spotted to a sample plate followed by 1 µl of matrix and allowed to air-dry. The matrix used was a 20 mg/ml solution of 2,4,6-trihydroxyacetophenone in 50% acetonitrile containing 0.1% trifluoroacetic acid and 0.05% (w/v) dodecylmaltoside. Samples were analyzed in the linear, positive ion mode using a delayed extraction of 300 ns and a grid voltage of 87.8%, and they were subject to a 25 kV accelerating voltage. External calibrations were performed routinely using horse apomyoglobin and bovine serum albumin (16,951 and 66,430 Da, respectively). Data were routinely processed using the 19-point Savitsky-Golay smoothing option included in the software provided by the manufacturer.


    Acknowledgments
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
The authors thank Janet A. Weigel for iodination of streptavidin and Leona Medved for assistance with the manuscript. This research was supported by National Institute of General Medical Sciences grant GM35978 from the National Institutes of Health.


    Abbreviations
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Biotin-PEO-maleimide, (+)-biotinyl-3-maleimido-propionamidyl-3,6-dioxaoctanediamine; HA, hyaluronan or hyaluronic acid; HAS, HA synthase; MALDI-TOF MS, matrix-assisted laser desorption ionization time-of-flight mass spectrometry; NEM, N-ethylmaleimide; NTA, nitrilotriacetic acid; ORF, open reading frame; SDS–PAGE, sodium dodecyl sulfate–polyacrylamide gel electrophoresis; seHAS, Streptococcus equisimilis HA synthase; spHAS, Streptococcus pyognes HA synthase; suHAS, Streptococcus ubris HA synthase.


    Footnotes
 
1 To whom correspondence should be addressed Back


    References
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Abatangelo, G. and Weigel, P.H. (eds) (2000) New frontiers in medical sciences: redefining hyaluronan. Elsevier Science B.V., Amsterdam.

Bliss, J.M. and Silver, R.P. (1996) Coating the surface: a model for expression of capsular polysialic acid in Escherichia coli K1. Mol. Microbiol., 21, 221–231.[ISI][Medline]

DeAngelis, P.L. (1999) Hyaluronan synthases: fascinating glycosyltransferases from vertebrates, bacterial pathogens, and algal viruses. Cell. Mol. Life Sci., 56, 670–682.[ISI]

DeAngelis, P.L., Papaconstantinou, J., and Weigel, P.H. (1993a) Isolation of a Streptococcus pyogenes gene locus that directs hyaluronan biosynthesis in acapsular mutants and in heterologous bacteria. J. Biol. Chem., 268, 14568–14571.[Abstract/Free Full Text]

DeAngelis, P.L., Papaconstantinou, J., and Weigel, P.H. (1993b) Molecular cloning, identification, and sequence of the hyaluronan synthase gene from Group A Streptococcus pyogenes. J. Biol. Chem., 268, 19181–19184.[Abstract/Free Full Text]

DeAngelis, P.L., Jing, W., Drake, R.R., and Achyuthan, A.M. (1998) Identification and molecular cloning of a unique hyaluronan synthase from Pasteurella multocida. J. Biol. Chem., 273, 8454–8458.[Abstract/Free Full Text]

DeAngelis, P.L., Jing, W., Graves, M.V., Burbank, D.E., and Van Etten, J.L. (1997) Hyaluronan synthase of chlorella virus PBCV-1. Science, 278, 1800–1803.[Abstract/Free Full Text]

Evered, D. and Whelan, J. (eds) (1989) The biology of hyaluronan. Ciba Fnd. Symp., 143, 1–288.

Fenderson, B.A., Stamenkovic, I., and Aruffo, A. (1993) Localization of hyaluronan in mouse embryos during implantation, gastrulation and organogenesis. Differentiation, 54, 85–98.[ISI][Medline]

Frillingos, S., Sahin-Toh, M., Wu, J., and Kaback, H.R. (1998) Cys-scanning mutagenesis: a novel approach to structure-function relationships in polytopic membrane proteins. FASEB J., 12, 1281–1299.[Abstract/Free Full Text]

Heldermon, C.D., DeAngelis, P.L., and Weigel P.H. (2001) Topological organization of the hyaluronan synthase from Streptococcus pyogenes. J. Biol. Chem., 276, 2037–2046.[Abstract/Free Full Text]

Hill, A.V. (1913) The combinations of haemoglobin with oxygen and with carbon monoxide. Biochem. J., 7, 471–480.

Itano, N. and Kimata, K. (1996a) Expression cloning and molecular characterization of HAS protein, a eukaryotic hyaluronan synthase. J. Biol. Chem., 271, 9875–9878.[Abstract/Free Full Text]

Itano, N. and Kimata, K. (1996b) Molecular cloning of human hyaluronan synthase. Biochem. Biophys. Res. Commun., 222, 816–820.[ISI][Medline]

Jung, K., Jung, H., and Kaback, H.R. (1994) Dynamics of lactose permease of Escherichia coli determined by site-directed fluorescence labeling. Biochemistry, 33, 3980–3985.[ISI][Medline]

Knudson, C.B. and Knudson, W. (1993) Hyaluronan-binding proteins in development, tissue homeostasis, and disease. FASEB J., 7, 1233–1241.[Abstract/Free Full Text]

Kumari, K. and Weigel, P.H. (1997) Molecular cloning, expression, and characterization of the authentic hyaluronan synthase from Group C Streptococcus equisimilis. J. Biol. Chem., 272, 32539–32546.[Abstract/Free Full Text]

Laemmli, U.K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature, 227, 680–685.[ISI][Medline]

Laurent, T.C. and Fraser, J.R.E. (1992) Hyaluronan. FASEB J., 6, 2397–2404.[Abstract/Free Full Text]

Lin, Y., Mahan, K., Lathrop, W.F., Myles, D.G., and Primakoff, P. (1994) A hyaluronidase activity of the sperm plasma membrane protein PH-20 enables sperm to penetrate the cumulus cell layer surrounding the egg. J. Cell Biol., 125, 1157–1163.[Abstract]

Moxon, E.R. and Kroll, J.S. (1990) The role of bacterial polysaccharide capsules as virulence factors. Curr. Top. Microbiol. Immunol., 6, 65–85.

Shyjan, A.M., Heldin, P., Butcher, E.C., Yoshino, T., and Briskin, M.J. (1996) Functional cloning of the cDNA for a human hyaluronan synthase. J. Biol. Chem., 271, 23395–23399.[Abstract/Free Full Text]

Spicer, A.P., Augustine, M.L., and McDonald, J.A. (1996) Molecular cloning and characterization of a putative mouse hyaluronan synthase. J. Biol. Chem., 271, 23400-23406.[Abstract/Free Full Text]

Tlapak-Simmons, V.L., Baggenstoss, B.A., Clyne, T., and Weigel, P.H. (1999a) Purification and lipid dependence of the recombinant hyaluronan synthases from Streptococcus pyogenes and Streptococcus equisimilis. J. Biol. Chem., 274, 4239–4245.[Abstract/Free Full Text]

Tlapak-Simmons, V.L., Baggenstoss, B.A., Kumari, K., Heldermon, C., and Weigel, P.H. (1999b) Kinetic characterization of the recombinant hyaluronan synthases from Streptococcus pyogenes and Streptococcus equisimilis. J. Biol. Chem., 274, 4246–4253.[Abstract/Free Full Text]

Tlapak-Simmons, V.L., Kempner, E.S., Baggenstoss, B.A., and Weigel, P.H. (1998) The active streptococcal hyaluronan synthases (HASs) contain a single HAS monomer and multiple cardiolipin molecules. J. Biol. Chem., 273, 26100–26109.[Abstract/Free Full Text]

Toole, B.P. (1997) Hyaluronan in morphogenesis. J. Intern. Med., 242, 35–40.[ISI][Medline]

Towbin, H., Steahelin, T., and Gordon, J. (1979) Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications. Proc. Natl Acad. Sci. USA, 76, 4350–4354.[Abstract]

Voss, J., Hubbell, W.L., Hernandez-Borrell, J., and Kaback, H.R. (1997) Site-directed spin-labeling of transmembrane domain VII and the 4B1 antibody epitope in the lactose permease of Escherichia coli. Biochemistry, 36, 15061–15066.

Wang, Q. and Kaback, H.R. (1999) Location of helix III in the lactose permease of Escherichia coli as determined by site-directed cross-linking. Biochemistry, 38, 16777–16782.[ISI][Medline]

Ward, P.N., Field, T.R., Ditcham, W.G., Maguin, E., and Leigh, J.A. (2001) Identification and disruption of two discrete loci encoding hyaluronic acid capsule biosynthesis genes hasA, hasB and hasC in Streptococcus uberis. Infect. Immun., 69, 392–399.[Abstract/Free Full Text]

Watanabe, K. and Yamaguchi, Y. (1996) Molecular identification of a putative human hyaluronan synthase. J. Biol. Chem., 271, 22945–22948.[Abstract/Free Full Text]

Weigel, P.H. (1998) Bacterial hyaluronan synthases. In Hascall, V.C., and Yanagishita, M., (eds), Science of hyaluronan today. Available online at: www.GlycoForum.gr.ip.

Weigel, P.H., Hascall, V.C., and Tammi, M. (1997) Hyaluronan synthases. J. Biol. Chem., 272, 13997–14000.[Free Full Text]

Wessels, M.R., Goldberg, J.B., Moses, A.E., and Dicesare, T.J. (1994) Effects on virulence of mutations in a locus essential for hyaluronic acid capsule expression in Group A streptococci. Infect. Immun., 62, 433–441.[Abstract]

Wessels, M.R., Moses, A.E., Goldberg, J.B., and Dicesare T.J. (1991) Hyaluronic acid capsule is a virulence factor for mucoid Group A streptococci. Proc. Natl Acad. Sci. USA, 88, 8317–8321.[Abstract]

Whitnack, E., Bisno, A.L., and Beachey, E.H. (1981) Hyaluronate capsule prevents attachment of Group A streptococci to mouse peritoneal macrophages. Infect. Immun., 31, 985–991.[ISI][Medline]