Heparanases: endoglycosidases that degrade heparan sulfate proteoglycans (original) (raw)

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Division of Molecular Biology and Biochemistry, School of Biological Sciences, University of Missouri-Kansas City, Kansas City, MO 64110, USA

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Abstract

Heparanases are endoglycosidases that cleave the heparan sulfate glycosaminoglycans from proteoglycan core proteins and degrade them to small oligosaccharides. Inside cells, these enzymes are important for the normal catabolism of heparan sulfate proteoglycans (HSPGs), generating glycosaminoglycan fragments that are then transported to lysosomes and completely degraded. When secreted, heparanases are thought to degrade basement membrane HSPGs at sites of injury or inflammation, allowing extravasion of immune cells into nonvascular spaces and releasing factors that regulate cell proliferation and angiogenesis. Heparanases have been described in a wide variety of tissues and cells, but because of difficulties in developing simple assays to follow activity, very little has been known about enzyme diversity until recently. Within the last 10 years, heparanases have been purified from platelets, placenta, and Chinese hamster ovary cells. Characterization of the enzymes suggests there may be a family of heparanase proteins with different substrate specificities and potential functions.

Accepted on March 19, 2001;

Introduction

Interactions between the extracellular environment and the cell surface influence cell proliferation, differentiation, migration, and shape. Heparan sulfate proteoglycans (HSPGs) play an important role in these interactions, because they are components of both basement and plasma membranes (Iozzo et al., 1994; Bernfield et al., 1999). The anionic heparan sulfate (HS) glycosaminoglycan chains bind to extracellular matrix and cell surface proteins, providing a framework for matrix organization and cell–cell or cell–matrix interactions. However, HSPGs play more than just a structural role. Both basement membrane and cell surface HSPGs bind a wide variety of protein ligands that are involved in wound repair, morphogenesis, host defenses, and lipid metabolism (Conrad, 1998; Bernfield et al., 1999). Numerous studies indicate that the protein–HS interactions are functionally important. Association of the ligand with HS glycosaminoglycans may activate or stabilize the ligand (Pillarisetti et al., 1997; Conrad, 1998; Lyon and Gallagher, 1998; Woods et al., 1998; Sperinde and Nugent, 1998) or be involved in directing the molecule to a different intracellular or extracellular location (Sperinde and Nugent, 1998; Tumova et al., 1999; Mahley and Ji, 1999). Therefore, to understand how these physiological processes are controlled, it is important to determine how the interactions between ligands and HSPGs are regulated.

Much attention has been focused on the sequence of the HS glycosaminoglycan to which ligands bind. HS chains are originally synthesized as a polysaccharide of alternating N-acetylglucosamine (GlcNAc) and glucuronic acid residues (GlcUA). In the Golgi, a series of enzymatic reactions occur that replace acetyl groups with sulfate groups, epimerize the glucuronic acid to iduronic acid (IdoUA), and add sulfate to the C-6 and C-3 hydroxyl groups of glucosamine and the C-2 hydroxyl groups of uronic acid residues (Lindahl et al., 1998). Because these modification reactions are incomplete, the final HS molecule has a domain structure (Lyon and Gallagher, 1998): N- and O-sulfate groups are clustered in IdoUA-rich sequences (S-domains or NS domains), which are separated by regions of [GlcNAc-GlcUA] disaccharide repeats (NA domains) that contain very little O-sulfate (Figure 1). Bridging these two domains are “mixed sequences” (or NA/NS domains) where GlcNAc disaccharides and GlcNS disaccharides alternate. In most HS species, S-domains range from 2 to 9 disaccharides and are separated by mixed and unmodified sequences that average 16 to 18 disaccharides. Ligands bind to the S-domains and mixed sequences (Conrad, 1998; Lyon and Gallagher, 1998); in some cases the ligand binds to a specific arrangement of sulfated glucosamine and uronic acid residues, and in others the interaction is primarily electrostatic. In either case, the formation of these S-domain sequences will determine whether a ligand can bind to the HSPG. Therefore, one way to regulate the interaction of a ligand with extracellular or cell surface proteoglycans is to regulate the synthesis of specific S-domains. Indeed, the ability of specific ligands to bind HSPGs can change on cell differentiation or age, due to differences in the fine structure of the S-domains (Brickman et al., 1998; Feyzi et al., 1998; Molist et al., 1998).

Another way to regulate the interaction of cell surface and matrix HSPGs with protein ligands is to degrade the HS glycosaminoglycans. This is done through the action of extracellular and intracellular endoglycosidases called heparanases. Heparanases can remove all binding sites from the proteoglycan by cleaving the HS chain from the core protein, or they can destroy specific ligand binding sites by cleaving within S-domains and/or mixed sequences. In addition to removing binding sequences, heparanases have been proposed to have other functions. Extracellular heparanases are believed to play roles in remodeling basement membranes after injury or at inflammation sites (Hoogewerf et al., 1995; Parish et al., 1998; Ihrcke et al., 1998; Dempsey et al., 2000) and in regulating cell growth and differentiation by releasing growth factors that are bound to extracellular HSPGs (Ishai-Michaeli et al., 1990). Heparanases in endosomes (Brauker and Wang, 1987; Yanagishita and Hascall, 1992; Tumova et al., 1999) are responsible for degrading internalized cell-associated HSPGs. In addition to generating short chain substrates for the lysosomal exoglycosidases (Kresse and Glössl, 1987), the intracellular heparanases may release bound ligands from their proteoglycan receptors once the complex is internalized (Tumova et al., 1999) or modulate the modification state of cell surface HSPGs by removing the HS chains from core proteins that recycle through the Golgi back to the plasma membrane (Edgren et al., 1997). The short HS oligosaccharides generated by either extracellular or intracellular heparanases may regulate the binding of ligands to cell surface or extracellular HSPGs, stabilize the ligand (Moscatelli, 1988; Pillarisetti et al., 1997), or facilitate the transport of the ligand to other sites of action (Sperinde and Nugent, 1998; Tumova et al., 1999). Degradation of HSPGs by heparanases may also be an important mechanism to prevent proteoglycans or long HS glycosaminoglycans from associating with molecules that would be deleterious to the cell, such as the β-amyloid-HSPG complex deposited in senile plaques of Alzheimer’s disease (Snow et al., 1994; Bame et al., 1997).

Pitfalls and challenges of assaying heparanase activity

The role of heparanases in regulating proteoglycan function has not received as much attention as the synthesis of specific binding sequences on HS glycosaminoglycans. One reason for this is the difficulties in designing an assay that could easily monitor the degradation of HS chains so the enzymes could be purified and characterized. Unlike the bacterial polysaccharide lyases that cleave HS or heparin chains by an eliminase mechanism and generate an unsaturated product (Linhardt et al., 1986), the mammalian heparanases are hydrolases, so activity cannot be monitored spectrophotometrically. The heparanase substrate is also difficult to obtain, because most of the HS molecules purified from tissues have already been endoglycosidically processed. Some researchers have used heparin (Oosta et al., 1982), but because it is more highly modified and lacks the domain structure of HS (Lyon and Gallagher, 1998), it may not be physiologically relevant. Over the past 25 years a number of different heparanase assays have been developed using radioactive HS substrates. They include monitoring the size of the glycosaminoglycan by gel filtration chromatography or gel electrophoresis (Höök et al., 1975; Klein and von Figura, 1979; Nakajima et al., 1984; Marchetti et al., 1997), precipitation of long HS chains under conditions where short oligosaccharides are soluble (Oldberg et al., 1980; Graham and Underwood, 1996; Tumova and Bame, 1997), release of radioactivity from immobilized HS or heparin (Oosta et al., 1982; Hoogewerf et al., 1995; Gonzalez-Stawinski et al., 1999) or from 35S-HSPG incorporated into a basement membrane (Matzner et al., 1985), and the inability of the heparanase product to be retained on a HS-ligand column (Freeman and Parish, 1998). One problem with all of these assays is that they may underestimate the number of functional heparanases in cells. Assays that measure enzyme activity by the monitoring the sizes of the substrates and products may not be able to discriminate between heparanases with different substrate specificities. Alternatively, assays that measure the release of HS chains from basement membranes or rely on the loss of ligand binding may miss activities that are not able to access the matrix glycosaminoglycans or do not affect ligand binding sites. There is another potential problem that is unique to the assay that measures the release of HS chains from basement membranes. Because it is likely that secreted heparanases will be incorporated into the matrix due to their ability to bind to HSPGs, one may be assaying an activator of a heparanase present in the matrix substrate, rather than the heparanase itself.

Research on heparanases has also been slow because the protein is not very abundant; estimates from a variety of sources indicate the enzyme(s) are less than 0.0001% of the total cellular protein (Oosta et al., 1982; Freeman and Parish, 1998; Bame et al., 1998; Gonzalez-Stawinski et al., 1999). Therefore, getting enough material to purify adequate enzyme for characterization has been a challenge. The stability of the enzymatic activity is also a problem; the earliest purification studies reported loss of enzymatic activity after one or two chromatographic steps (Klein and von Figura, 1979; Oldberg et al., 1980). Later work has been able to maintain activity throughout the steps needed to purify the protein(s) (Oosta et al., 1982; Hoogewerf et al., 1995; Goshen et al., 1996; Freeman and Parish, 1998; Bame et al., 1998), although in our experience, activity is most stable in a partially purified form (Bame, unpublished data).

By the mid-1990s the variety of assays had firmly established that heparanase activity was present in a wide variety of tissues and cells (Höök et al., 1975; Klein and von Figura, 1979; Oldberg et al., 1980; Matzner et al., 1985; Sewell et al., 1989; Lider et al., 1990; Bame, 1993), including tumor cells (Ogren and Lindahl, 1975; Nakajima et al., 1984; Peretz et al., 1990; Laskov et al., 1991). The activities were shown to be β-endoglucuronidases, cleaving both HS and heparin to shorter oligosaccharides with a GlcNAc–GlcUA sequence at the newly formed reducing end (Klein and von Figura, 1979; Oldberg et al., 1980; Oosta et al., 1982; Nakajima et al., 1984; Bame and Robson, 1997). The glycosaminoglycan substrate had to have N-sulfate and free carboxyl groups, and be larger than 3000 Da to be cleaved (Klein and von Figura, 1979; Oldberg et al., 1980; Irimura et al., 1986). However, it was not known how many proteins were responsible for the described heparanase activities. Only Oosta et al. (1982) had succeeded in purifying a 130-kDa heparanase protein from platelets, but no amino acid sequence of the purified enzyme was reported. In the past six years, however, great strides have been made in purifying and/or characterizing heparanases from platelets, placenta, and Chinese hamster ovary (CHO) cells.

Heparanase proteins

CTAP-III

Hoogewerfand colleagues reported the purification of a heparanase from platelets in 1995 (Hoogewerf et al., 1995). Unlike the earlier enzyme (Oosta et al., 1982), activity was associated with a 9–10-kDa protein derived from the CXC-chemokine, platelet basic protein. Platelet basic protein is proteolytically processed to three smaller chemokines, connective tissue activating peptide-III (CTAP-III), β-thromboglobulin, and neutrophil activating peptide-2, that function in inflammation, wound healing, and growth regulation (Proudfoot et al., 1997). Because these functions also involve HSPGs (Conrad, 1998; Bernfield et al., 1999), the finding that the chemokine could degrade the glycosaminoglycan suggested a novel mechanism to regulate HSPG activities. The idea that CTAP-III was a heparanase was initially questioned, because some commercial preparations of the chemokine lacked activity (Graham and Underwood, 1996). However, later studies showed commercially prepared β-thromboglobulin could cleave an HS octasaccharide (Pikas et al., 1998). Genetic studies finally established that CTAP-III does indeed encode a heparanase. Recombinant human CTAP-III was fused to a 17-kDa cellulose binding domain and expressed in Escherichia coli (Rechter et al., 1999). The bacterially produced protein was shown to have heparanase activity because it degraded soluble 35S-HSPGs and released 35S-HS from basement membranes. An antibody against CTAP-III indicates its presence in T cells, PMN lymphocytes, and placental extracts (Rechter et al., 1999), suggesting that the secretion of CTAP-III may play a role in facilitating the movement and migration of cells through extravascular tissues.

Hpa1 heparanase

The next reports of a purified heparanase were of a 45–50-kDa glycoprotein found in placenta (Goshen et al., 1996) and platelets (Freeman and Parish, 1998; Gonzalez-Stawinski et al., 1999). Although of similar size, it was unclear whether the placenta and platelet heparanases were the same protein, because different assays and schemes were used to purify the activity and only one of the reports gave any protein sequence (Gonzalez-Stawinski et al., 1999). Another question was whether this smaller heparanase was similar to the 130-kDa enzyme originally purified from platelets (Oosta et al., 1982). One group found that the purified platelet heparanase could form higher molecular weight aggregates (Gonzalez-Stawinski et al., 1999), suggesting a possible explanation for the original observation of a larger protein. Once the heparanase was cloned (Vlodavsky et al., 1999; Hulett et al., 1999; Kussie et al., 1999; Toyoshima and Nakajima, 1999) and functionally expressed in insect cells (Vlodavsky et al., 1999) and COS-7 cells (Hulett et al., 1999), it became clear that the platelet and placenta proteins were identical. This novel heparanase (designated Hpa1) is initially synthesized as an inactive 65-kDa glycoprotein that is cleaved at the N-terminus to generate the active enzyme (Vlodavsky et al., 1999; Hulett et al., 1999). It is not yet known whether the activation reaction occurs inside cells or after the enzyme is secreted. Expression of the full-length cDNA is required to generate active heparanase (Hulett et al., 1999), indicating that the N-terminus of the proheparanase is essential for the expression or function of the enzyme. A recent report suggests that the active Hpa1 heparanase may actually be a heterodimer, where the 50-kDa protein acts in concert with a tightly associated, noncovalently linked 8-kDa peptide, derived from the N-terminus of the 65-kDa proheparanase (Fairbanks et al., 1999). Hpa1 heparanase homologues are found in bovine and rat, however, there is no comparable protein in Drosophila melanogaster or Caenorhabditis elegans, suggesting that the endoglycosidic cleavage of HSPGs in these organisms (Bernfield et al., 1999; Yamada et al., 1999) is carried out by a different enzyme.

Northern blot analysis shows that Hpa1 is highly expressed in placenta and lymphoid organs (Hulett et al., 1999; Kussie et al., 1999; McKenzie et al., 2000), suggesting that the primary functions of this enzyme are to assist in leukocyte migration, remodel basement membranes at sites of inflammation, and regulate processes involved in embryo implantation and pregnancy (Vlodavsky et al., 1999; Hulett et al., 1999; Dempsey et al., 2000). Studies with the purified Hpa1 heparanase show that at physiological pH the enzyme binds to the extracellular matrix or cell surface HSPG but is inactive (Gilat et al., 1995; Ihrcke et al., 1998). It has been proposed that when the pH is lowered, which would occur at sites of inflammation or matrix damage, the bound enzyme becomes active and cleaves the HSPGs it is bound to (Gilat et al., 1995; Ihrcke et al., 1998; Dempsey et al., 2000). Antibodies made against Hpa1 heparanase show the enzyme is localized to the subendothelium (Dempsey et al., 2000), supporting the hypothesis that the heparanase is already present at sites where HSPG degradation may occur. Hpa1 heparanase is also highly expressed in tumors and metastatic cell lines derived from a variety of tissues (Vlodavsky et al., 1999; Hulett et al., 1999; Kussie et al., 1999; Dempsey et al., 2000; McKenzie et al., 2000). Indeed, when the hpa1 cDNA is transfected into cells they acquire a highly metastatic phenotype (Vlodavsky et al., 1999), suggesting a direct role of Hpa1 heparanase in cancer processes, such as metastasis.

Hpa2 heparanase

Using the Hpa1 heparanase amino acid sequence in a BLAST search of a proprietary human EST database, another putative heparanase gene, Hpa2, has been recently identified that is 35% identical to the Hpa1 protein (McKenzie et al., 2000). Due to alternative splicing, this gene may encode three putative heparanase proteins that range in size from 48 to 60 kDa. However, as of yet, it is not known whether any of these proteins have enzymatic activity. The different tissue distribution of hpa2 mRNA and hpa1 mRNA (McKenzie et al., 2000) suggests that if they are enzymes, the Hpa2 heparanases may have different functions. Hpa2 is expressed in human brain, mammary gland, prostate, small intestine, testis, and uterus (McKenzie et al., 2000). As with Hpa1 heparanase, there are no Hpa2 heparanase homologues found in the D. melanogaster and C. elegans genomes.

CHO cell heparanases

Neither hpa1 nor hpa2 mRNA is expressed well in heart, kidney, liver, lung, skeletal muscle, thymus, and thyroid (McKenzie et al., 2000). Because these tissues are known to express cell surface HSPGs, this finding suggests there may be another heparanase enzyme that catalyzes the endoglycosidic cleavage of the internalized HS glycosaminoglycans. CHO cells have been used as the source of heparanases responsible for degrading endocytosed cell surface HSPGs. CHO cell heparanase activities do not appear to be secreted, and cellular fractionation studies indicate that the activities reside in endosomes (Tumova et al., 1999). In addition, the hpa1 transcript has not been detected in purified CHO cell mRNA (Bame and Venkatesan, unpublished data), suggesting that, if present, the Hpa1 heparanase is expressed at very low levels in CHO cells and may not be the primary activity that cleaves the cell-associated HSPGs.

Characterization of the short HS glycosaminoglycans purified from CHO cells suggests the presence of multiple intracellular heparanases, because some short HS products had an S-domain at the nonreducing end, whereas others had the S-domain at the reducing end of the molecule (Bame and Robson, 1997). Four heparanase activities with different molecular properties have been purified from CHO cell homogenates (Bame et al., 1998). C1A heparanase is a 37–48-kDa protein that elutes from a cation exchange column between 0.4 and 0.8 M NaCl and can be separated from the 30-kDa C1B heparanase protein by gel filtration chromatography. The other two heparanase subtypes, C2A and C2B, are ∼ 45-kDa proteins that elute from the cation exchange column between 0.9 and 1.2 M NaCl and can be separated by DNA-cellulose chromatography.

At present, only the most abundant CHO activity, C1A heparanase, has been further characterized (Bame and Venkatesan, unpublished data). The enzyme has been purified from rat liver as well, indicating that the expression of this heparanase is not restricted to CHO cells. Unlike the Hpa1 heparanase, the C1A heparanase does not appear to be a glycoprotein because the activity does not bind to a ConA column. Preliminary sequencing and antibody studies indicate the ∼ 40-kDa C1A heparanase contains the N-terminal domain of the ∼ 80-kDa proteins ezrin, radixin, and moesin (ERM), suggesting that the enzyme may be derived from these molecules. The ERM proteins are involved in regulating cell morphology and adhesion (Tsukita and Yonemura, 1999), and although they are known as heparin-binding proteins (Hirao et al., 1996), none of them has been reported to have heparanase activity. The hypothesis that processed forms of ERM proteins are intracellular heparanases is an attractive one, since the widespread distribution of at least one ERM protein across species (Tsukita and Yonemura, 1999) and in cells (Doi et al., 1999) would ensure that every internalized HSPG would be endoglycosidically cleaved. However, further work needs to be done to establish whether C1A heparanase is derived from the ERM proteins or if it is a newly identified member of the family of proteins that contain this N-terminal associated domain (Chishti et al., 1998). Also, the other CHO heparanase subtype proteins need to be characterized to see if they are related to the C1A or Hpa enzymes, or if they are additional unique members of a family of heparanases.

Heparanase substrate specificity

CTAP-III

CTAP-III heparanase cleaves both HS and heparin glycosaminoglycans to short oligosaccharides and is active between pH 5.0 and 7.5, with optimal activity at pH 5.8 (Hoogewerf et al., 1995). Initial characterization of the activity suggested that CTAP-III was a novel heparanase because it cleaved the HS glycosaminoglycan at glucosamine residues (Hoogewerf et al., 1995). However, it has been subsequently shown that, like the other described heparanase activities, CTAP-III is a β-endoglucuronidase (Pikas et al., 1998). It is not known whether CTAP-III heparanase requires a specific sequence or modification to recognize and cleave the HS chain, although it is clear that the glycosaminoglycan substrate must be O-sulfated (Pikas et al., 1998).

Hpa 1 heparanase

Depending on the assay, Hpa1 heparanase is active from pH 4.0 to 7.5, with maximal activity at pH 5.5 to 5.8 (Gilat et al., 1995; Graham and Underwood, 1996; Freeman and Parish, 1998). It cleaves both HS and heparin to ∼ 5-kDa oligosaccharides (Freeman and Parish, 1998; Pikas et al., 1998), suggesting the enzyme does not rely on the HS domain structure (Figure 1) to recognize the substrate. Instead, evidence indicates Hpa1 heparanase recognizes a specific chain modification to bind and cleave the HS glycosaminoglycan. Clues to the glycosaminoglycan sequence recognized by Hpa1 heparanase came from studies of the enzyme expressed in mouse and human melanoma cell lines (Marchetti et al., 1997). If HS chains are digested with melanoma Hpa1 heparanase, the shorter oligosaccharides are no longer able to bind a peptide derived from heparin-interacting protein (HIP). Conversely, if the HIP peptide is included in the in vitro assay, Hpa1 heparanase is unable to cleave the HS chains. The inhibition of melanoma Hpa1 is specific for the HIP peptide, because comparable amounts of another HS ligand, basic fibroblast growth factor (bFGF), have no affect on Hpa1 heparanase activity (Marchetti et al., 1997). These results suggest that Hpa1 heparanase and HIP recognize the same glycosaminoglycan sequence. Because HIP was shown to compete with antithrombin III for binding to HS glycosaminoglycans (Liu et al., 1997), it appears that Hpa1 heparanase may cleave the chain either within or adjacent to the pentasaccharide sequence that binds the serpin (Figure 2). This finding is supported by studies that show Hpa1 heparanase, purified from platelets or hepatoma cells, can cleave an octasaccharide that contains the highly sulfated antithrombin III binding sequence (Thunberg et al., 1982; Pikas et al., 1998). Interestingly, if the 2-O-sulfate group on the IdoUA residue adjacent to the Hpa1 cleavage site is removed (Figure 2), the octasaccharide is no longer a substrate (Pikas et al., 1998). Based on this result, it has been proposed that a 2-O-sulfate modification is essential for Hpa1 heparanase to recognize and cleave the HS chain. However, it is likely that the sequence context of the 2-O-sulfate group will be important for Hpa1 heparanase recognition as well, because bFGF, which does not affect Hpa1 heparanase activity, also binds to a sequence that contains 2-O-sulfated IdoUA residues (Figure 2).

A primary function of Hpa1 heparanase is to degrade basement membrane HSPGs at sites of injury or inflammation (Vlodavsky et al., 1999; Hulett et al., 1999; Dempsey et al., 2000). This will disrupt the basement membrane to allow extravasion of inflammatory cells and liberate growth factors and chemokines that induce proliferation and migration of endothelial cells and fibroblasts. It is likely that only a small portion of the basement membrane proteoglycan HS chains are accessible to the heparanase, because a variety of matrix proteins, cell adhesion proteins, and growth factors will be bound to sites on the glycosaminoglycan (Iozzo et al., 1994). By recognizing just a specific sequence or modification, Hpa1 heparanase does not need to rely on other structural features of the glycosaminoglycan that may be obscured when the proteoglycan is incorporated in the matrix. It may also be important that anticoagulant activities at the site be inhibited, thus the observation that Hpa1 heparanase can cleave the HS chain so that the binding sites for antithrombin III (Marchetti et al., 1997; Pikas et al., 1998) and histidine-rich glycoprotein (Freeman and Parish, 1998) are destroyed may have functional significance (Conrad, 1998; Borza and Morgan, 1998). Genetic studies in which the gene is ablated will be quite revealing.

CHO cell C1A heparanase

All four CHO cell heparanases, including C1A, are active from pH 3.5 to 5.5and cleave long HS glycosaminoglycans to short chains of 6 to 8 kDa (Bame et al., 1998). However, rather than a specific modification or sequence, the CHO heparanases appear to use the HS domain structure (Figure 1) to recognize and cleave the glycosaminoglycan. None of the individual CHO heparanase subtypes degrade HS glycosaminoglycans in vitro if bFGF is included in the assay (Tumova and Bame, 1997; Bame et al., 1998). The inhibition of CHO heparanase activity is specific for bFGF, and a direct interaction between the HS chain and the growth factor at multiple sites is required to prevent degradation (Tumova and Bame, 1997). However, because the short heparanase products still contain S-domains that bind bFGF (Tumova and Bame, 1997), these findings indicte that the CHO enzymes are not cleaving the chain within this sequence. Instead they suggest that the heparanases use differences in sulfate content between the S-domains and unmodified sequences to orient themselves on the glycosaminoglycan and then cleave the chain in the mixed sequences (Figure 1; Bame and Robson, 1997). This hypothesis is supported by studies using undersulfated HS glycosaminoglycans synthesized by the CHO cell mutant, _pgs_E-606. This mutant is deficient in the modification enzyme N-deacetylase/N-sulfotransferase I (Bame and Esko, 1989; Aikawa and Esko, 1999), which determines the extent of sulfation and uronic acid epimerization of the HS glycosaminoglycan (Lindahl et al., 1998). The mutant cells synthesize nascent HS chains that have fewer S-domains than the wild-type glycosaminoglycan, although the spacing of the S-domains on mutant and wild-type chains are similar (Bame et al., 2000). When incubated with different CHO heparanase subtypes, only the portion of the _pgs_E-606 HS that is structurally similar to wild type is degraded (Bame et al., 2000). Characterization of the longer products from the _pgs_E-606 HS substrates suggests they are resistant to further degradation by the enzymes because they do not have internal S-domains (Bame et al., 2000).

The S-domains recognized by bFGF contain 2-O-sulfated IdoUA residues (Figure 2), however, this modification does not appear to be required for recognition by the CHO heparanases based on experiments using HS substrates that lack 2-O-sulfate groups. One of these substrates is the HS synthesized by the CHO cell mutant, _pgs_F-17, which is deficient in 2-O-sulfotransferase activity (Bai and Esko, 1996). A partially purified preparation of CHO heparanases can degrade the _pgs_F-17 HS, although not as efficiently or to the same extent as wild-type glycosaminogycans (Bai et al., 1997). Structural studies of the mutant glycosaminoglycan show that in addition to the lack of 2-O-sulfate groups, the HS chains have an increased number of GlcNS residues (Bai and Esko, 1996), which should affect the domain structure of the molecule by shortening the unmodified regions (Figure 1). Therefore, instead of the absence of 2-O-sulfate groups, it may be that the decreased size of the unsulfated regions affects the ability of the CHO enzymes to recognize and cleave the mutant _pgs_F-17 HS chains. The other HS substrate that lacks 2-O-sulfate groups is synthesized by a mouse that had the 2-O-sulfotransferase gene inactivated by a targeted gene-trap technique (Merry et al., unpublished data). Although the HS from the mutant mouse does not have 2-O-sulfate groups, the polysaccharide is sulfated similarly to the wild-type glycosaminoglycan, due to a slight increase in the length of S-domains and an increase in 6-O-sulfate groups within the S-domains (Merry et al., unpublished data). When 3H-HS isolated from either wild-type or 2-O-sulfotransferase-mutant mice were incubated exhaustively with purified C1A heparanase, both glycosaminoglycans were completely cleaved to chains of 3 to 5-kDa (Bame, unpublished data). This result shows that 2-O-sulfate groups are not required for C1A heparanase to recognize and degrade the substrate.

One function of intracellular heparanases is to degrade the long HS glycosaminoglycans to shorter chains that are the substrates for the lysosomal exoglycosidases. Unlike the HSPGs in basement membranes, the endocytosed HS chains should be relatively free of bound ligands, so it is possible that rather than a specific glycosaminoglycan sequence, the intracellular heparanases recognize the substrate using a three-dimensional structure formed by regularly spaced S-domains. Studies with heparin oligosaccharides show that the conformation of the IdoUA residues in S-domains cause both the 2-O-sulfate and 6-O-sulfate groups to be clustered on one side of the polysaccharide (Mulloy et al., 1994; Mulloy and Forster, 2000). This structure would allow the heparanase protein to differentiate easily between sulfated and nonsulfated regions of the chain and bind the glycosaminoglycan appropriately. Another advantage of a three-dimensional recognition model for the intracellular heparanases is that the secondary structure formed by the IdoUA-rich sequences does not change significantly when the placement of O-sulfate groups within the S-domain is altered (Mulloy et al., 1994). Thus, even if the fine structure of the S-domains changes on developmental state or aging (Brickman et al., 1998; Feyzi et al., 1998; Molist et al., 1998), the heparanase should still be able to degrade the glycosaminoglycan and generate the short HS fragments for lysosomal catabolism or other cellular functions.

Conclusions

In the past several years it has been established that the degradation of HSPGs in mammals is catalyzed by at least two different heparanase enzymes, CTAP-III and Hpa1 heparanase. Other studies support the idea that there are additional heparanases with different physical properties and substrate specificities that are also responsible for degrading HSPGs. In addition to confirming that the C1A and Hpa2 proteins are indeed heparanases, future studies need to elucidate the functional role of the individual enzymes in vivo. Because both CTAP-III and Hpa1 heparanase have been proposed to degrade basement membrane HSPGs are they redundant activities or do they have unique functions that depend on the cells that express them? Could the low level of hpa1 mRNA observed in liver or kidney (McKenzie et al., 2000) generate enough enzyme to account for the intracellular degradation of HSPGs in these tissues, or is another enzyme, such as C1A heparanase essential for this process? How are the activities of the secreted and intracellular heparanases regulated in vivo? How do the different enzymes bind the HS substrate and catalyze the cleavage reaction? Are there homologues for the mammalian heparanases in Drosophila or C. elegans? Answers to these questions will not only provide valuable information on how heparanases degrade HSPGs and thereby regulate HSPG functions, but they will also give clues for developing compounds that can selectively inhibit the heparanase activities associated with inflammation or tumor metastasis, yet protect the activities essential for normal cellular functions.

Abbreviations

bFGF, basic fibroblast growth factor; CHO, Chinese hamster ovary; CTAP-III, connective tissue activating peptide-III; ERM, ezrin, radixin, and moesin; GlcNAc, N-acetylglucosamine; GlcUA, glucuronic acid; HIP, heparin-interacting protein; HS, heparan sulfate; HSPG, heparan sulfate proteoglycans; IdoUA, iduronic acid

Fig. 1. Heparan sulfate glycosaminoglycan structure and potential heparanase cleavage sites. Evidence suggests that Hpa1 heparanase cleaves a sequence within the highly modified S-domain (indicated by open arrow; Marchetti et al., 1997; Pikas et al., 1998), and studies with C1A heparanase suggest the enzyme cleaves within the mixed sequences (indicated by closed arrow; Bame and Robson, 1997; Bame et al., 2000).

Fig. 2. Heparan sulfate sequences. (a) A heptasaccharide containing the pentasaccharide sequence recognized by antithrombin III (indicated by brackets; Pikas et al., 1998). When this sequence was part of an octasaccharide, the Hpa1 heparanase cleaved the molecule at the position indicated by the arrow. If the 2-O-sulfate group was removed from the IdoUA, the octasaccharide was no longer cleaved (Pikas et al., 1998). (b) The minimal pentasaccharide sequence recognized by basic fibroblast growth factor (indicated by brackets; Lyon and Gallagher, 1998). If bFGF is included in the enzyme assay, Hpa1 heparanase is not inhibited (Marchetti et al., 1997), but C1A heparanase is (Bame et al., 1998), suggesting that the two enzymes recognize the HS substrate differently. The open arrow indicates a potential cleavage site for C1A heparanase.

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