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Department of Basic Sciences and Aquatic Medicine, Norwegian School of Veterinary Science, PO Box 8146 Dep., N-0033 Oslo, Norway
Correspondence
Grethe Skretting
grethe.skretting{at}medisin.uio.no
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Present address: Hematological Research Laboratory, Ullevål University Hospital, N-0407 Oslo, Norway. 
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). This change in protein structure results in heavy accumulation of PrPSc in the central nervous system (CNS), particularly at the terminal stage of the disease (Caughey et al., 2001). Normally, PrP mRNA is expressed widely in tissues, particularly in neurons (Caughey et al., 1988; Brown et al., 1990; Cagampang et al., 1999). The importance of PrP in prion diseases has been shown in studies using PrP-knockout mice. In the CNS, PrP expression in neurons is required for the development of neuropathology (Brandner et al., 1996), and in peripheral tissues, expression of PrP is important for efficient transmission of infectivity from the periphery to the CNS (Blättler et al., 1997). Studies in transgenic mice overexpressing PrP have shown that the pace and route of infection are influenced by the expression levels of the host-encoded PrPC (Blättler et al., 1997; Glatzel & Aguzzi, 2000).
Uptake of the infectious agent from the alimentary tract is considered the natural route of infection in TSEs (Hadlow et al., 1982). Pathogenesis studies have shown that the gut-associated lymphoid tissue (GALT) is the first site of accumulation for PrPSc in scrapie in sheep and mice (Kimberlin & Walker, 1989; Andréoletti et al., 2000; Heggebø et al., 2000). The major component of the GALT in young sheep is a continuous aggregation of lymphoid follicles termed the ileal Peyer's patch (PP) (Landsverk et al., 1991). During its life, the ileal PP is responsible for generating the vast majority of B lymphocytes in the circulation and peripheral lymphoid tissues (Reynolds & Morris, 1983; Gerber et al., 1986; Press et al., 1996) and for diversification of the pre-immune antibody repertoire (Reynaud et al., 1991; Lucier et al., 1998). This lymphoid organ involutes with age and is reduced dramatically by the age of 18 months. The follicles of the ileal PP consist predominantly of B lymphocytes supported by an extensive network of mesenchymal stromal cells including follicular dendritic cells (FDCs) and reticular cells, along with a population of tingible body macrophages and a few T lymphocytes (Halleraker et al., 1990; Nicander et al., 1991; Press et al., 1992). Most studies in scrapie-infected mice demonstrate that FDCs are the dominant cell type harbouring PrPSc in the lymphoreticular system (McBride et al., 1992; Brown et al., 1999; Mabbott et al., 2000). It has been shown that FDCs may enhance the spread of the infectious agent through surface accumulation of PrPSc (Shlomchik et al., 2001). However, there is still discussion as to whether the FDCs in germinal centres are the cells of the lymphoreticular system that sustain conversion of PrPC to PrPSc in TSEs (Shlomchik et al., 2001; Prinz et al., 2002).
At present, little is known about the molecular mechanisms involved in the uptake of the infectious agent from the gut lumen or its accumulation in lymphoid follicles. Indirect evidence for the involvement of GALT in sheep has been derived from oral challenge of lambs with the infectious agent. Heggebø et al. (2000) reported that PrP levels detected by immunohistochemistry were increased in follicles of the ileal PP as early as 1 week after challenge, whereas the presence of PrPSc was only detectable 5 weeks after challenge (Heggebø et al., 2003a). These investigators and others (Andréoletti et al., 2000; Heggebø et al., 2000, 2003a) have speculated that uptake occurs across the follicle-associated epithelium (FAE) overlying the dome of the ileal PP, a proposition supported by experimental studies using cultured M-cell monolayers (Heppner et al., 2001). However, both M-cell-dependent and -independent routes for the transport of PrPSc across the epithelium have been proposed (Ghosh, 2004; Huang & MacPherson, 2004). Both routes imply that, after uptake across the epithelium, PrPSc is transported to lymphoid tissues, possibly by migrating populations of dendritic cells (DCs) (Bruce et al., 2000; Huang et al., 2002). Others have suggested that neuroendocrine cell types, via their expression of PrPC, might play an important role in the internalization of PrPSc from gut lumen (Marcos et al., 2004).
More knowledge on PrP gene expression in GALT will provide an insight into how and where the infectious agent is taken up from the gut lumen and how the infection proceeds before it enters the CNS via the enteric nervous system (ENS) in the gut wall or by other pathways. It is, however, important to note that the site where PrPC is produced might not be the same as the site where it is converted to PrPSc. Similarly, the site of conversion could be different from the eventual site of PrPSc accumulation.
The aim of the present work was to compare the expression of PrP mRNA with the presence of PrPC in the GALT of lambs at an age where they are most susceptible to scrapie and consequently most likely to be infected, using a combination of quantitative and distributional techniques. We present the first quantification of PrP mRNA distribution in tissue compartments of the ileal PP by using laser-capture microdissection and real-time RT-PCR. These results are supported by in situ hybridization and immunohistochemistry.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; St Rose et al., 2006). Immediately after killing, tissue pieces from ileal PP were removed from the animals and placed with the mucosal surface down on liver slices to protect the mucosa during freezing. Slices of cerebellum, 5 mm in thickness, were also collected. These tissues were snap-frozen in 1,1,1-trifluorethane/1,1,1,2 tetrafluorethane/pentafluorethane (R404A; Ausimont), chilled in liquid nitrogen and stored at –80 °C until use.
Laser-capture microdissection.
Sections of 14 µm thickness were cut by using a cryostat (Leitz Cryostat 1720) and mounted on special membrane slides for laser microdissection (Molecular Machines and Industries). The sections were air-dried at room temperature and stored at –80 °C until use. Prior to microdissection, the membrane sections were stained with RNase-free haematoxylin and dried at room temperature. Laser-capture microdissection of tissue sections was performed by using the SLµCut laser microdissection system (Molecular Machines and Industries). The SLµCut equipment is provided with an automated UV dissection system coupled to video imaging. Tissue samples were removed securely (Fig. 1a, b) by using an adhesive membrane, which protects the tissue on the slide against cross-contamination. To preclude internal variation, several pieces of each desired compartment were microdissected to obtain an area corresponding to 1x106 µm2 or approximately 500–1000 cells depending on the cell density, i.e. at least 10 different follicles from each animal. The microdissected compartments were follicle, dome, interfollicular area, outer submucosa (i.e. area of submucosa immediately adjacent to the inner muscular layer), muscular layer, FAE, lamina propria and villous epithelium (Fig. 1c).
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Quantitative real-time RT-PCR.
Quantitative real-time RT-PCR was performed by using a One-step qPCR core kit (Eurogentec). Primers were designed to span across intron sections by using PrimerExpress 1.5 (Applied Biosystems). The expression level was measured by relative quantification using glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as the reference gene. GAPDH has been shown to be expressed more stably across different tissues than other frequently used housekeeping genes (Foss et al., 1998; Garcia-Crespo et al., 2005). Each quantification target was amplified in triplicate and a control lacking the template for each mastermix was always included in the experiments.
Primers and hybridization probes used for the quantitative real-time RT-PCR are as follows: Ovis aries PrP: forward, 5'-TCCCAGAGACACAGATCCAACTT-3'; reverse, 5'-GATCCAACTGCCTATGTGGCTT-3'; probe, 5'-FAM-ACCATGATGACTTCTATCTGCTGTGATTCAGCT-TAMRA-3'. Ovis aries GAPDH: forward, 5'-TGATTCCACCCATGGCAAGT-3'; reverse, 5'-CCACGTACTCAGCACCAGCAT-3'; probe, 5'-FAM-TCCACGGCACAGTCAAGGCAGAGAA-TAMRA-3'. Real-time RT-PCR was carried out in an ABI PRISM 7700 (Applied Biosystems) using the following uniform temperature profile: 30 min at 48 °C (reverse transcription), then 10 min at 95 °C (denaturation), followed by 40 cycles of 30 s at 95 °C, 15 s at 56 °C and 60 s at 60 °C. The same cycling profile was used for all real-time RT-PCRs. The data were analysed by using the Sequence Detection System v1.9.1 (Applied Biosystems). Statistical differences between the different tissue compartments were evaluated by using a paired t-test and the distribution of all datasets was tested by using the Kolmogorov & Smirnov test. Differences in expression between compartments were considered to be significant with values of probability (P) <0.05 (InStat; GraphPad Software).
In situ hybridization.
Frozen sections (12 µm) were cut with a cryostat (Leitz Cryostat 1720) and mounted on positively charged slides (Superfrost Plus; Menzel-Gläser). Frozen-tissue sections from the same lambs as were used for quantitative real-time RT-PCR were selected. The sections were air-dried and stored at –80 °C until use. To increase the sensitivity, a cocktail of two digoxigenin (DIG)-labelled cRNA nucleotide fragments of the coding and the 3'-untranslated regions of PRNP mRNA was utilized, each covering approximately 700 bp of the ORF or the 3' UTR. PCR fragments containing T7 and SP6 promoters were prepared from plasmid clones of the two regions by using vector-derived M13 forward and reverse primers. The amplified PCR products were gel-purified and used as templates for synthesis of the DIG-labelled cRNA antisense and sense probes by using T7 or SP6 polymerases (Roche) following the manufacturer's instructions. All RNA products were checked by gel electrophoresis and stored at –80 °C until use. Probe concentrations were determined by using spot tests. In situ hybridization was carried out according to the method of Barthel & Raymond (1993) with some modifications. Briefly, the tissue sections were rehydrated, fixed in 4 % paraformaldehyde in PBS, treated with 10 µg proteinase K ml–1 (Sigma-Aldrich) for 5 min at 37 °C followed by fixation in 4 % paraformaldehyde in PBS, then treated with acetic anhydride and dehydrated. Approximately 100 µl hybridization mixture containing 100 ng of each DIG-labelled RNA probe was applied directly to each air-dried section and the sections were incubated in a humidity chamber with coverslips for 15 h at 60 °C. Non-specific binding of probes was removed by digestion for 30 min at 37 °C with 20 µg RNase A ml–1 (Sigma-Aldrich). The probes were detected by using an anti-DIG antibody coupled to alkaline phosphatase. Labelling was visualized by using NBT/BCIP as substrate. For each of the analysed tissue sections, sense probes were applied to serial sections as a negative control to confirm the specificity of the hybridization.
Immunolabelling for PrPC on frozen sections.
Frozen sections (10 µm) were placed on positively charged slides (Superfrost Plus; Menzel-Gläser), allowed to dry for 1 h and fixed in 10 % formol/calcium for 15 min. The following monoclonal antibodies (mAbs) of the IgG1 isotype were used to detect PrP: L42, P4 (both kindly provided by Dr Martin Groschup, Greifswald-Insel Reims, Germany) and 6H4 (Prionics). In the following steps, a TSA-indirect kit (NEN Life Science Products) was used according to the manufacturer's instructions with some modifications. All incubations were performed in a humid chamber at room temperature and sections were washed in PBS with 0.05 % Tween 20 between incubations unless otherwise stated. To block non-specific binding of antibodies and binding of streptavidin in the TSA kit to endogenous biotin, sections were incubated in avidin (Vector Laboratories) diluted 1 : 6 in TNB blocking buffer [0.1 M Tris/HCl (pH 7.5), 0.15 M NaCl, 0.5 % blocking reagent supplied in the TSA kit] for 30 min. After gentle removal of the blocking solution, sections were incubated overnight at 4 °C with primary antibody diluted in biotin (Vector Laboratories) : TNB 1 : 6. To detect binding of primary antibodies, the sections were incubated in biotinylated sheep anti-mouse Ig (RPN 1001; Amersham Biosciences) followed by a streptavidin–horseradish peroxidase (HRP) complex (P0397; DakoCytomation). Signal enhancement was performed by using biotinyl–thyramide followed by streptavidin–HRP; both reagents were supplied in the TSA kit. Finally, signals were developed with aminoethylcarbazole (AEC) as substrate. As negative-control sections, primary antibodies were replaced by an irrelevant antibody of the same isotype (IgG1) (BD PharMingen) or dilution buffer. Microphotographs were captured by using a Spot RT Slider digital camera (Diagnostic Instruments Inc.) mounted on a Leica DMRXA microscope (Leica Microsystems Wetzlar GmbH).
Detection of FDCs by immunohistochemistry.
Frozen sections (8 µm) were placed on positively charged slides (Superfrost Plus; Menzel-Gläser), allowed to dry for 1 h and fixed in 4 % formaldehyde for 2 min followed by 70 % ethanol for 10 min. Anti-FDC mAb CNA.42 (M7157; DakoCytomation) was used to detect an antigen expressed predominantly by FDCs in various species including sheep (Raymond et al., 1997; Lezmi et al., 2001). Sections were labelled by using a polymeric labelling kit (UltraVision LPValue detection system; Lab Vision Corporation) according to the manufacturer's instructions. Sections were incubated overnight at 4 °C with the CNA.42 antibody diluted 1 : 100 in TBS [0.1 M Tris/HCl (pH 7.5), 0.15 M NaCl] containing 1 % BSA. Labelling was detected by using AEC from the kit. Control sections were incubated overnight at 37 °C in neuraminidase solution [neuraminidase (N5631-1UN; Sigma-Aldrich) at 0.01 U ml–1 in 50 mM sodium acetate, 154 mM NaCl, 9 mM CaCl2, pH 5.5] before immunolabelling, as the CNA.42 reactive epitope is known to be destroyed by neuraminidase (Raymond et al., 1997).
Detection of FDCs by 5'-nucleotidase histochemistry.
Enzyme activity for 5'-nucleotidase was examined by using the method described by Müller-Hermelink et al. (1974) with minor modifications. In brief, frozen sections (7 µm) were air-dried at room temperature for 1–2 h and fixed for 5 min at 4 °C in 4 % formaldehyde containing 68 mM CaCl2 adjusted to pH 7.2 with NaOH. Reactivity for 5'-nucleotidase was detected by incubation with a solution composed of 1 mM adenosine 5'-monophosphate, 200 mM Tris/maleate buffer (pH 7.4), 1.3 mM Pb(NO3)2, 100 mM MgSO4.7H2O and 96 mM sucrose. The sections were incubated for 40 min at 37 °C, followed by treatment with a 0.2 % (NH4)2S solution for 1 min. The addition of 100 mM NaF to the incubation solution inhibited the reaction.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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) were laser-microdissected. PrP mRNA expression was detected in all examined compartments of the ileal PP, with distinct differences in the expression levels between the compartments (Fig. 2). Lowest expression was seen in the follicles and the villous epithelium. The dome, interfollicular area, FAE and the lamina propria showed a two- to threefold higher expression level compared with the level in the follicles. The highest expression level of PrP mRNA was found in the outer submucosa, and muscular layer. In these compartments, the levels were approximately five- to tenfold higher than in the follicles. All animals in this experiment displayed the same pattern regarding the distribution and levels of expression, and no major differences in the expression level in these samples could be seen between the sheep with different PrP genotypes. Pair-specific comparisons of expression levels in the different dissected tissue compartments are presented in Table 1.
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). Although PrP mRNA expression was detected in the follicles, villous and T-cell areas by quantitative real-time RT-PCR, labelling was not observed in these tissue compartments by using in situ hybridization (data not shown). As brain tissue is known to express high amounts of PrP mRNA (Tichopad et al., 2003; Ning et al., 2005), the cerebellum was used as a positive-control tissue to ensure that the probes were specific. In cerebellum, strong labelling was present in Purkinje cells and some scattered cells of the molecular and granule-cell layers (data not shown), which is in agreement with previous reports in sheep and human (Kubosaki et al., 2000; McLennan et al., 2001). In negative-control sections, in which the antisense PrP probe was replaced with a sense probe, no labelling was observed in any sections from the ileal PP or the cerebellum. When each of the two antisense RNA probes was applied separately, they both produced positive PrP mRNA-specific labelling (data not shown).
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). The distribution of PrP in the lamina propria and interfollicular area suggested the labelling of nerves or smooth-muscle cells (Fig. 4b). Labelling was demonstrated also in some large, mononuclear cells in these areas. Most follicles showed little or no labelling. In negative-control sections, no labelling caused by endogenous peroxidase or non-specific interactions caused by the IgG1 isotype-control antibody were observed (data not shown).
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) and there was strong staining for 5'-nucleotidase in the light central zone of the lymphoid follicles (Fig. 5a). A similar distribution of immunohistochemical labelling was observed with the CNA.42 antibody (Fig. 5b). This distribution of staining, obtained by using both of these methods, indicated the presence of FDCs in the light central zones of all follicles in all animals examined, and that at least 80 % of the follicles in a section seemed to be cut through the light central zone. No 5'-nucleotidase staining was detected in NaF-inhibited sections. In neuraminidase-treated sections, labelling with the CNA.42 antibody was abolished.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Heggebø et al., 2000). Given the assumption that potential sites of PrPSc conversion should display high levels of expression of PrPC (McBride et al., 1992), it was unexpected to find that the follicles showed very low levels of PrP mRNA. Indeed, the lymphoid follicles had the lowest level of expression of all compartments examined (Fig. 2). In mice, it has previously been reported that PrP mRNA levels may not correlate with PrPC levels in cells of the CNS (Ford et al., 2002a). Our results indicate, however, that the low levels of PrP mRNA in the follicles of sheep ileal PP were supported by the absence of labelling for PrPC by immunohistochemistry.
The expression of PrPC by FDCs has been reported to be critical for the conversion and accumulation of PrPSc in lymphoid follicles (Brown et al., 1999) and subsequent neuroinvasion in mouse scrapie (Mabbott et al., 2000). Similarly, the accumulation of PrPSc in sheep scrapie was found to be associated with FDCs in lymphoid follicles early in infection (Jeffrey et al., 2000; Heggebø et al., 2002). However, the expression level of PrP in FDCs has previously not been investigated in sheep. In cattle, FDCs isolated from lymphoid tissues were found to react only with one mAb against PrP, 6H4, indicating that a special isoform of PrPC was present in this type of cell (Thielen et al., 2001). In sheep, even though both 5'-nucleotidase enzyme activity and immunohistochemistry confirmed the presence of FDCs, they were not labelled by using 6H4 or two other mAbs in immunohistochemistry on frozen sections, suggesting a possible difference in the amount of PrPC present in FDCs between cattle and sheep.
The observed low levels of PrP expression in lymphoid follicles and FDCs of the ileal PP of sheep suggest that the PrPSc accumulation found in these sites during scrapie infection (Jeffrey et al., 2000; Heggebø et al., 2002) was not derived from conversion of PrPC produced by FDCs. It is more likely that preformed PrPSc is transported into the follicles and subsequently trapped on FDCs via receptors such as the complement receptor (Mabbott et al., 2001). Another possibility is that the levels of PrPC in FDCs are below the detection limits of the techniques used in the present study, and that both the conversion and accumulation of PrPSc still might take place in FDCs during the prolonged incubation period of scrapie. A third possibility, not investigated in this study, is that PrP expression may be increased in infected animals.
It is possible that the presence of DCs in the dome and the interfollicular regions can account for the significantly higher levels of PrP expression that were present in these compartments compared with the levels detected in lymphoid follicles (Kelsall & Strober, 1996; Huang & MacPherson, 2004; Defaweux et al., 2005). DCs are known to express PrPC (Burthem et al., 2001; Sugaya et al., 2002) and they form a dense layer of cells in the dome in the PP just beneath the FAE, where they are in close contact with M cells (Halleraker et al., 1990; Press et al., 1992; Kelsall & Strober, 1996). It has previously been shown that DCs can acquire PrPSc in vitro and that a small subpopulation of migrating DCs is able to take up and transport PrPSc from the gut lumen through the lymphatics to the lymphoid tissue (Huang et al., 2002). PrPSc accumulation in the dome and subsequently in central FDCs of follicles in the early phases of scrapie has been reported (Andréoletti et al., 2000). These accumulation patterns indicate that conversion of PrPSc might take place in the dome, a tissue compartment with higher levels of PrP mRNA expression levels than the follicles, before the misfolded protein is transported to and accumulates in association with FDCs of the follicles.
The highest levels of PrP mRNA expression were observed in the outer submucosa and the muscular layer and were probably the result of the high PrPC expression in the submucosal (Meissner's) and myenteric (Auerbach's) plexi of the ENS, as shown in mice (Ford et al., 2002b) and sheep (Heggebø et al., 2003b). The high expression levels that were detected by real-time RT-PCR in samples microdissected from the outer submucosa and the muscular layer were supported by in situ hybridization and immunohistochemistry. Both techniques revealed extensive labelling in neuronal cell bodies and satellite cells of the myenteric and submucosal nervous plexi, structures with substantial accumulation of PrPSc during scrapie (van Keulen et al., 1999; Heggebø et al., 2002). In the small intestine of mice, specific labelling of enteroendocrine cells and small enterocytes in the epithelium or myofibroblasts in the lamina propria has been reported (Ford et al., 2002b). Additionally, González et al. (2005) showed a faint and inconsistent FDC-like pattern of PrP immunolabelling in the lymphoid follicles. Labelling of these cell populations was not detected in the present work. This discrepancy may be due to the sensitivity of the in situ hybridization and immunohistochemistry protocols used in our study. It should be noted that species differences may account for varying levels of PrP expression, as may the differing functions of the ileal PP as opposed to conventional mucosal lymphoid tissue represented by jejunal PP in sheep and PP in mice (Reynolds & Morris, 1983; Landsverk et al., 1991).
To our knowledge, the present study is the first report on the expression levels and distribution pattern of PrP mRNA in the ileal PP in sheep. The finding that the follicles displayed the lowest levels of expression of PrP mRNA in the ileal PP focused attention on the relationship between PrPC expression, PrPSc conversion and accumulation of PrPSc. The molecular mechanisms by which PrPSc is taken up and transferred to nerves, and to what extent FDCs, M cells and DCs are involved in these processes, await further clarification, in particular the demonstration that the expression of PrPC is involved in the process of transfer of PrPSc from the gut lumen to subepithelial tissues. The refinement of laser-dissection techniques and an improvement in sensitivity of in situ hybridization protocols, along with accompanying studies on experimentally challenged scrapie animals, may enhance our understanding of these fundamental events in the peripheral pathogenesis of scrapie in sheep.
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ACKNOWLEDGEMENTS |
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Received 6 March 2006;
accepted 24 July 2006.
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