Departments of 2Surgery, 3Immunology, and 4Pediatrics, Mayo Clinic, Rochester, MN 55905, USA
Received on July 6, 1999; revised on November 16, 1999; accepted on November 16, 1999.
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Abstract |
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Key words: heparanase/heparan sulfate/hyperacute rejection/placenta/xenografts
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Introduction |
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Besides contributing to tissue homeostasis and development, heparan sulfate metabolism may be linked to disease. During inflammation, the extravasation of leukocytes is linked to loss of heparan sulfate from endothelial cells and extracellular matrices (Naparstek et al., 1984; Matzner et al., 1985
; Fridman et al., 1987
; Shimada and Ozawa, 1987
; Ishai-Michaeli et al., 1990
). Tumor cell metastatic potential also correlates with the ability to degrade heparan sulfate proteoglycans (Nakajima et al., 1988
; Vlodavsky et al., 1992
). Heparanase inhibitors dramatically reduce the incidence of lung metastases following injection of metastatic tumor cells in mice (Nakajima et al., 1983
; Vlodavsky et al., 1988
; Villanueva et al., 1989
). Tumor angiogenesis may be promoted by releasing heparan sulfate-associated growth factors sequestered in the underlying extracellular matrix, amplifying the effect of heparan sulfate degradation (Vlodavsky et al., 1988
; Weiss et al., 1988
; Ishai-Michaeli et al., 1990
). Immune cell interactions can be bolstered in vitro by heparan sulfate as shown by increasing T cell proliferation and cytotoxicity and by stimulating antigen-presenting cells (Wrenshall et al., 1991
, 1994).
Heparan sulfate proteoglycans might be shed from cells and tissues by one or more of four mechanisms: (1) proteolytic cleavage of the protein core, (2) phospholipase-mediated cleavage of a lipid anchor, (3) release of peripheral membrane proteoglycans by disruption of ionic interactions, and (4) cleavage of glycosaminoglycan chains by endoglycosidases (Brunner et al., 1991; Bernfield et al., 1992
; Yanagishita, 1992
; Yanagishita and Hascall, 1992
; Ihrcke and Platt, 1996
). Of these, glycosaminoglycan cleavage would be expected to have the most profound impact on tissue physiology, since it is predominantly the glycosaminoglycan chains that confer biological function to heparan sulfate proteoglycans (Salmivirta et al., 1996
; Stringer and Gallagher, 1997
; Lindahl et al., 1998
). Activated platelets can release heparanase at sites of inflammation (Oosta et al., 1982
). Because heparan sulfate plays a critical role in maintaining vascular integrity and in development, we postulated that elaboration of heparanase might be tightly regulated. Platelet heparanase is inactive at physiological pH, but becomes highly active at acidic pH, as occurs during inflammation (Gilat et al., 1995
; Ihrcke et al., 1998
). However, heparanase retains its ability to bind to heparan sulfate molecules at physiological pH (Ihrcke et al., 1998
) and is, possibly, stored in extracellular domains.
Heparanase activity has been variously attributed to proteins of molecular mass ranging between 4060 kDa (Freeman and Parish, 1998; Gonzalez-Stawinski et al., 1999
; Goshen et al., 1996
; Graham and Underwood, 1996
; Sandbäck Pikas et al., 1998
), to a 9 kDa CXC chemokine CTAP-III (Hoogewerf et al., 1995
), or a 134 kDa endoglycosidase activity derived from platelets (Oosta et al., 1982
). The discrepancies regarding the identity of heparanase and the difficulty in obtaining pure heparanase preparations has precluded detailed mechanistic studies on the regulation of heparanase expression and activity. In this report, we have begun to characterize the contribution of heparanase in health and disease by identification of human heparanase cDNAs and the gene encoding human heparanase, and by localization of heparanase protein expression in normal tissues and in diseased organs.
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Results |
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The predicted heparanase amino acid sequence revealed several relevant features that are shown in Figure 1. BLAST searches of the protein database or translated nucleic acid databases failed to reveal any known protein of similar sequence. Hileman et al. (1998) and Cardin and Weintraub (1989)
identified heparin- or heparan sulfate-binding protein consensus motifs XBBBXXBX and XBBXBX, thought to be important for ionic interactions with glycosaminoglycan ligands. Human heparanase contains two regions of clustered basic amino acids, 272278 (PRRKTAKM) and 157162 (QKKFKN), which conform to these binding motif patterns (Figure 1). The highly basic peptide KRRKLRV, amino acids 426432, is also present in heparanase protein. Platelets release heparanase as a 55 kDa soluble enzyme, suggesting that protein processing may occur to produce the mature enzyme. Hulett et al. (1999)
reported processing of the precursor heparanase between amino acids 157158 to give the mature product. Six putative N-glycosylation sites are present in the heparanase conceptual sequence. We have shown, previously, that purified heparanase binds to Concanavalin A (Gonzalez-Stawinski et al., 1999
), suggesting that one or more glycosidation sites may be used in vivo.
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We have postulated that inflammatory conditions induce both the release of platelet heparanase and activation of heparanase catalytic activity by lowering the pH of the vascular microenvironment (Ihrcke et al., 1998). To examine this hypothesis in vivo, we analyzed porcine hearts undergoing hyperacute rejection for platelet aggregation, heparanase expression, and loss of vascular heparan sulfate. Blood vessels in a normal porcine heart are devoid of platelet aggregates, as evidenced by the lack of immunostaining with anti-human CD9, which recognizes primate platelets but no pig structures. Heparanase was observed with the subendothelial smooth muscle layer surrounding blood arteries in normal hearts (Figure 6A), but heparanase was not found on the luminal blood vessel surface (inset, Figure 6A). Normal tissues contained abundant heparan sulfate deposition within extracellular matrices surrounding blood vessels, extending up to the endothelial cell surface (inset, Figure 6B). However, when we examined tissues from hyperacute rejecting hearts, dense platelet aggregates were present in blood vessel lumen (not shown) and focal heparanase deposition was found on endothelial cell surfaces (Figure 6C). We asked if loss of heparan sulfate was correlated with the presence of heparanase in inflamed tissues. Examination of adjacent tissue sections demonstrated heparanase deposition (Figure 6C inset) on the luminal face of the blood vessel, which was coincident with loss of endothelial cell heparan sulfate staining (Figure 6D, inset arrow). Close inspection of the blood vessel surface revealed several nucleated cells, which appear to be transversing the endothelial cell boundary (inset, Figure 6C). These observations are consistent with heparanase activation in vivo in hyperacute organ rejection, and that heparanase activity might contribute to loss of vascular integrity, leading to the demise of organ transplants.
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Discussion |
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The heparanase cDNA sequence we report is identical to the sequence recently reported by Vlodavsky et al. (1999) and Hulett et al. (1999)
; however, we report, here, differences in the tissue expression of heparanase protein that could impact on its physiological role(s). Vlodavsky et al. (1999)
report localization of heparanase protein in primary or metastatic tumor tissues where heparanase might contribute to the degradation of extracellular matrices and, thereby, contribute to the dissemination of the tumor cells. However, heparanase mRNA and protein were not detected in normal tissues. In contrast, Northern hybridization data, shown here, suggests normal human tissues do express low steady-state levels of the 1.8 kb transcript, which is upregulated in placental tissues where abundant heparanase activity is found. Increased steady-state levels of this 1.8 kb transcript were also found in several human cancer cell lines, as shown here and in lymphoid tissues, such as bone marrow and peripheral blood leukocytes (Hulett et al., 1999
). Under physiological conditions, heparanase activity may be limited to acidified membrane compartments, such as the lysosome, where heparan sulfate proteoglycans may be degraded to achieve steady-state levels. Under conditions of inflammation, heparanase may be secreted by platelets or leukocytes into local blood vessel lumen, compromising vascular integrity and allowing the passage of blood-borne cells into interstitial spaces. In addition, inactive forms of heparanase protein may be tethered to extracellular matrices. We report, here, finding heparanase protein in subendothelial locations in both normal tissues and tissues undergoing humoral-induced vascular rejection episodes. Antibodies against N-terminal peptides found in the precursor domain did not recognize heparanase protein within placental or heart tissues in immunofluorescence experiments, suggesting that the tethered heparanase molecules are not unprocessed precursors but might be mature protein (data not shown). We would postulate that heparanase may be activated in inflamed tissue as local pH decreases, as discussed below. The loss of endothelial cell heparan sulfate in the latter tissues, but not the former, is consistent with heparanase activation under inflammatory conditions.
The anti-heparanase antibodies used in our studies recognized peptide epitopes of the mature active protein, and were shown to specifically recognize denatured heparanase by Western immunoblots, and native platelet heparanase by ELISA. Moreover, by employing a stringent assay to measure inhibition of heparanase activity, we show the anti-heparanase antibodies recognize a significant fraction of active heparanase. To our knowledge, this is the first report describing antibody reagents that affect heparanase activity.
Given the detection of heparanase protein in normal tissues and the pathophysiologic consequences, were heparanase to be active, it would not be surprising that tight regulatory mechanisms control enzyme activity. We have postulated that the activity of expressed heparanase protein is regulated by microenvironmental conditions, particularly by pH. Heparanase binds to but does not degrade heparan sulfate glycosaminoglycans at physiological pH (Gilat et al., 1995; Ihrcke et al., 1998
). Heparanase endoglycosidase activity is optimal between pH 5.0 and 6.5; however, the enzyme is much less active when the pH is above 7.0 (Oosta et al., 1982
; Gilat et al., 1995
; Freeman and Parish, 1998
; Ihrcke et al., 1998
). Heparanase activity progressively decreases at neutral pH, but can be reactivated under acidic conditions (Ihrcke et al., 1998
). This property confers an important regulatory mechanism that may confine heparanase action to injured vascular sites, where inflammatory conditions lower the pH and are, thus, favorable for heparanase activity, thereby preventing systemic activation and inadvertent damage to distal endothelium. We have demonstrated, here, in pathophysiological findings of hyperacute rejection, where inflammatory vascular lesions abound, that heparanase becomes activated on endothelial surfaces with the concomitant loss of heparan sulfate and vascular integrity.
Additional mechanisms may exist to promote or inhibit heparanase activity in normal tissues and during development. Heparanase mRNA is expressed in many human tissues and immortalized tumor cell lines, but the pattern of expression differs, qualitatively, in tissues where abundant heparanase activity has been reported, suggesting posttranscriptional regulatory mechanisms act upon heparanase mRNA stability. The most abundant heparanase transcript found in steady-state mRNA populations appears to be too small to encode the mature protein product and lacks methionine initiation codons in the proper open reading frame. Regulation of heparanase activity may also involve competing protein interactions by blocking enzyme access to its substrate, as may occur by the cell surface protein heparan sulfate/heparin-interacting protein (HIP), first identified in uterine epithelial cells and, subsequently, found to be expressed in various epithelial and endothelial cells (Liu et al., 1997). HIP binds to the same oligosaccharide sequence as antithrombin III (Liu et al., 1997
) and antagonizes heparan sulfate digestion by heparanase, presumably by competing for the same binding recognition site in the glycosaminoglycan chain (Marchetti et al., 1997
). Expression of competing heparan sulfate-binding proteins may, in part, explain why abundant heparanase proteins are present in placental tissues, but adverse pathophysiological episodes normally do not occur during gestation. However, this notion raises the possibility that some complications in pregnancy, such as defective placenta formation in preeclampsia or abnormal maternal bleeding episodes, could be, in part, attributed to defective heparanase regulation.
The development of molecular tools, as described here, to study heparanase expression and regulation will no doubt contribute to our understanding of how heparanase functions during development, in tissue homeostasis and in disease. The ability to overexpress recombinant heparanase proteins will allow detailed investigation of the structural basis governing the pH-dependence of heparanase catalytic activity. Such knowledge can lead to the rational design of molecules, which may block structural transitions or sterically-hinder the heparanase catalytic active site or ligand-binding domain, thereby precluding heparanase digestion of heparan sulfate, and may be beneficial as a therapeutic intervention in disease states.
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Materials and methods |
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Cell culture
The human CHRF-288 megakaryoblast cell line, (Fugman et al., 1990), was cultured at 37°C in Fischers medium (Life Technologies, Grand Island, NY) supplemented with 10% fetal bovine serum (FBS), 100 U/ml penicillin-streptomycin, 2 mM L-glutamine, and 1 mM sodium pyruvate (Life Technologies).
Heparanase activity assays
Heparanase activity was assayed using 3H-heparan sulfate-Sepharose beads, prepared as described previously (Ihrcke et al., 1998). Washed platelets were resuspended in heparanase assay buffer (0.1 M sodium acetate, pH 5.0, 0.1 mg/ml bovine serum albumin, 0.01% Triton X-100; and protease inhibitors 0.5 mM phenylmethylsulfonyl fluoride, 10 µg/ml leupeptin, and 10 µg/ml pepstatin A), and subjected to several freezethaw cycles (70°C/37°C). Clarified lysates were obtained by centrifugation at 34,000 x g for 30 min at 4°C. Platelet lysate, 100 µg protein, was added to 350 µl heparanase buffer, 50 µl 3H-heparan sulfate-Sepharose beads equilibrated in the same buffer, and incubated at 37°C for 4 h. Release of 3H-heparan sulfate was quantitated by measuring the radioactivity in supernatant fractions by scintillation counting. Assays were performed in triplicate and each experiment was performed at least twice on separate days. Activity is reported as the mean amount of radioactivity (c.p.m.) released per 100 µg protein. Protein concentrations were determined by the detergent-compatible Lowry protein assay (Bio-Rad, Hercules, CA) using BSA as standards.
Generation of anti-heparanase antibodies
Mice were immunized with a synthetic peptide conjugate (Bio-Synthesis, Lewisville, TX) KLRVYLHCTNTDN (amino acids 430442) found within human heparanase. Responding mice were sacrificed and their splenic B cells fused with FO-SP2/O myeloma cells (Galfré and Milstein, 1981). Hybridoma lines secreting anti-heparanase antibodies were identified by ELISA and cloned by limiting dilution. Isotype determination was performed using goat anti-mouse
-chain specific or goat anti-mouse µ-chain specific antibodies (Sigma Chemicals, St. Louis, MO) in ELISA reactions. Rabbits were immunized using the same peptide conjugate to generate polyclonal antibodies. Additional peptides CKYGSIPPDVEEK (amino acids 128140) and TKVLMASVQGSKRRK (amino acids 417431) were coupled to maleimide-activated keyhole limpet hemocyanin carrier protein (Pierce) and used to immunize mice, as described above. To demonstrate the specificity of the anti-heparanase monoclonal and polyclonal antibodies, competitive ELISA assays were performed with fixed concentrations of platelet extract and anti-heparanase antibody in PBS, 0.1% BSA with increasing amounts of the peptide KLRVYLHCTNTDN. Alkaline phosphataseconjugated goat
-mouse IgM or goat
-rabbit IgG (diluted 1:1000 in PBS, 0.1% BSA) were used to detect primary immune complexes, followed by incubation with disodium p-nitrophenyl phosphate (Sigma Chemicals). Antibody binding was quantitated by measuring the absorbance at 405 nm. Western immunoblotting was performed by resolving human platelet extracts on 1020% gradient denaturing polyacrylamide gels, followed by transfer to PVDF membranes. Goat
-rabbit IgG or rabbit
-mouse IgM alkaline phosphatase-conjugated antibodies were used to detect primary immune complexes with NBT/BCIP reagents. To test for the ability of antibodies to inhibit heparanase activity, human platelet extracts (100 µg) were incubated in the presence of anti-heparanase monoclonal antibodies (100 µg) for 3 h, at room temperature, under conditions which are permissible for catalytic activity, (0.1 M sodium acetate, pH 6.5, 0.1 mg/ml bovine serum albumin, 0.01% Triton X-100; and protease inhibitors). Aliquots were removed to tubes containing immobilized 3H-heparan sulfateSepharose beads and were incubated 1.5 h further in the presence of protein A/G agarose beads, to which goat anti-mouse µ-chain antibodies had been coupled. Following incubation, the beads were pelleted and the supernatant removed for assay of heparanase activity. Immunoglobulin isotype controls, 2C18, which recognizes human complement factor C5a, and mock-treated protein A/G beads, were tested likewise.
Chromosomal localization of the human heparanase gene
Fluorescence in situ hybridization mapping of human metaphase chromosomes was performed using digoxigenin dUTP-labeled DNA probes, which were combined with sheared human DNA and hybridized to normal metaphase chromosomes in a solution containing 50% formamide, 10% dextran sulfate, and 20x SSC. Specific hybridization signals were detected by incubating the hybridized slides in fluoresceinated anti-digoxigenin antibodies followed by counterstaining with DAPI.
Northern blot hybridization
Northern blots containing 2 µg poly(A)+ mRNA from normal human tissues or from a variety of human cancer cell lines (Clontech, Inc., Palo Alto, CA) were hybridized to a 475-bp 32P-labeled probe, derived from the 3' end of the human heparanase cDNA (coordinates 10041479), which was generated by PCR-amplification using primers Hep1, 5'-CTGGCAAGAAGGTCTGGTT-3', and Hep3, 5'-GACAGATTTGGAAAGTAATCC-3'. Probes were radiolabeled to a specific activity of 1.0 x 109 c.p.m./µg by random-hexamer primer extension (Boehringer Mannheim, Indianapolis, IN) and [-32P]dCTP (Amersham, Arlington Heights, IL). Hybridization was performed at 68°C in ExpressHyb solution (Clontech). Blots were washed twice in 2x SSC, 0.05% SDS at room temperature, then twice in 0.1x SSC, 0.1% SDS at 50°C for 30 min, and exposed to Kodak XAR x-ray film. The blots were subsequently stripped and reprobed with a human ß-actin probe for normalization of the amount of RNA loaded per lane. Blots were exposed to film for 90 h with an intensifying screen at 80°C to detect heparanase expression, whereas exposure times for detecting ß-actin were 3 h.
RNA preparation, RT-PCR, and 5'-RACE (rapid amplification of DNA ends)
Total RNA was prepared from CHRF-288 cells by the guanidinium thiocyanate method (Chomczynski and Sacchi, 1987). Heparanase cDNAs were synthesized using Hep3 anti-sense primer and AMV reverse transcriptase (Boehringer Mannheim) and 3'poly(dA)-tailed with dATP and TdT. Two rounds of nested PCR amplification were performed with Expand PCR polymerase (Boehringer Mannheim) using anti-sense primers Hep4, 5'-GATCGAAGTTTTCATCCAC-3', and Hep5, 5' GTGATGCCATGTAACTG-3', and oligo(dT) or an anchor primer provided by the manufacturer (Boehringer Mannheim). Alternatively, human placental cDNA (Clontech) was utilized as the starting material for nested PCR amplification and used high temperature "touchdown" PCR cycling conditions in a Perkin-Elmer GeneAmp9600 thermocycler (94°C, 30 sec; 5 cycles at 94°C for 5 sec, 72°C for 4 min; 5 cycles at 94°C for 5 sec, 70°C for 4 min; 25 cycles at 94°C for 5 sec, 68°C for 4 min). Heparanase clones were identified by producing a diagnostic PCR product of 235 bp using a Hep7 (5'-GGCACGAGGAGGCAATGAACCT-3'),-Hep5 primer pair. The nucleotide sequence of positive clones was determined by the dideoxynucleotide chain termination method.
Indirect immunofluorescence
Representative specimens of human placenta (obtained by caesarian section), normal porcine heart tissue, and porcine hearts undergoing hyperacute rejection after xenotransplantation into baboon recipients, were snap-frozen in precooled isopentane and stored at 85°C until cyrosectioning. Tissue sections (4 µm) were cut using a Leica CM 3050 cryostat and mounted on positively charged microscope slides, briefly heat-fixed, and stored at 85°C. Sections were acetone-fixed 10 min at 4°C, and postfixed for 2 min in Tris-buffered 1% paraformaldehyde, 1 mM EDTA, pH 7.2. Formalin-fixed, paraffin-embedded tissue sections were prepared by placing tissue specimens in 10% neutral-buffered formalin, pH 7.2, for 18 h at room temperature, after which samples were processed and embedded into paraffin blocks. Tissue sections (4 µm) were obtained using a rotary microtome (model 2155, Leica) and mounted onto positively charged microscope slides. Primary antibodies were diluted and applied to tissue sections including monoclonals Hep1.1A (mouse IgM anti-human heparanase), HepSS-1 (mouse IgM anti-heparan sulfate, Seikagaku, Tokyo), and BA-2 (mouse IgG anti-CD9). Secondary fluorochrome-labeled antibodies were used to detect binding by the primary antibodies. Affinity-purified, FITC-conjugated goat F(ab')2 anti-mouse IgM (1:100) or anti-mouse IgG (1:200; Cappel-ICN, Aurora, OH) was applied to the primary antibody-stained tissue sections. A third layer of FITC-conjugated rabbit F(ab')2 anti-goat IgG (Cappel-ICN), diluted 1:50, and rhodamine-conjugated rabbit anti-human basement membrane GBM50 (diluted 1:100) were applied to tissue samples. All fluorochrome antibodies were preabsorbed with human and porcine serum prior to use. Sections were counterstained/coverslipped with Vectashield-DAPI (Vector Laboratories, Burlington, CA) for detection of nuclei and stored in the dark at 4°C until microscopic evaluation using a fluorescence research microscope (Leica DMRD). Photographic images were obtained utilizing a CCD digital camera (SPOT II, Diagnostic Instruments, Sterling Heights, MI).
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Acknowledgments |
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Abbreviations |
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Footnotes |
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References |
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