1 Institut de Génétique Humaine, CNRS UPR1142, Montpellier, France, 2 Unit of Cell Biology, Department of Biochemistry and Physiology, Faculty of Biology, University of Barcelona, Barcelona, Spain and 3 Department of Nephrology, University Hospital Lapeyronie, Montpellier, France
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Abstract |
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Methods. Two forms of amyloidosis were studied: experimental amyloid A (AA) and clinical ß2-microglobulin amyloidosis. We studied kidney, liver, and spleen in a mouse model, and examined surgically obtained carpal deposits from dialysis patients. We used light and electron microscopy with immunogold labelling for anti-ß2-microglobulin and anti-AA protein antibodies.
Results. AA amyloid fibril accumulation was associated with membrane lesions in basal, cytoplasmic organelle (endoplasmic reticulum, mitochondria), and nuclear membranes. Amyloid fibrils from ß2-microglobulin amyloidosis were also closely associated with elastic fibres and endothelial basement membrane. We observed proliferation of endothelial cells as well as basement membrane enlargement and disruption.
Conclusions. Vascular abnormalities, including endothelial enlargement, basement membrane modifications, and vascular proliferation were associated with amyloidosis. Amyloid fibrils have a high avidity for elastic fibres and are able to contact and damage the basement membrane, the cell and intracellular organelle membranes, as well as the nuclear envelope, suggesting a toxic effect of amyloid fibrils on cells.
Keywords: amyloid fibril toxicity; amyloidosis; endothelial cells; experimental amyloidosis; ß2-microglobulin
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Introduction |
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Most studies examining the pathogenesis of dialysis-related amyloidosis have focused on ß2-microglobulin concentrations, biochemical modifications, or both [4]. However, serum ß2-microglobulin levels may not accurately predict the risk of appearance of dialysis-related amyloidosis [5], and none of the known modifications of ß2-microglobulin fully explains its ability to precipitate in amyloid fibrils [6]. Other investigations have focused on cell participation in the genesis of amyloid fibrils [79]. These studies found that the cells surrounding the amyloid deposits are mainly of monocyte-macrophage lineage [79]. In addition, recent evidence suggests that macrophages have mainly a degradative role in dialysis-related amyloidosis [reviewed in 10].
In the present study we analysed the participation of vessels and particularly vascular cells in this disease process. We analysed structural modifications of vessels and extracellular matrix with electron microscopy, and located amyloid proteins with coupling electron microscopy and immunogold labelling. We studied both experimental and human amyloidosis to identify common pathological features. Our findings demonstrate that there is a vascular involvement in both human and experimental amyloidosis, and that amyloid fibrils have an affinity for elastic fibres and cause membrane disruption in both forms of the disease.
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Subjects and methods |
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To assess the kinetics and organ involvement of experimental amyloidosis properly, mice were killed at 4 h, 1 day, 4 days and 1, 2, 3 and 4 weeks after the initial induction. The spleen, liver, and kidney were removed and examined for amyloid deposition.
Patients
Amyloid deposits were obtained from the carpal tunnel of three dialysis patients, aged 65±4 years with dialysis-related amyloidosis. Their renal diseases included two chronic glomerulonephritis cases and one autosomal dominant polycystic kidney disease. None of the patients had systemic diseases known to be associated with amyloidosis.
Sample preparation and immunohistochemistry
Congo red staining
Congo red, morphology studies, and immunogold methods were performed as previously described [11]. Unfixed amyloid deposits were immersed in OCT compound (Miles Inc., Diagnostics Division, Elkhart, USA), frozen and stored at -70°C. Cryostat sections of 10 µm were transferred onto gelatin-coated (0.3%) glass slides and stained with Congo red (Searle Scientific Services, High Wycombe Bucks, UK). Briefly, sections were dehydrated in 70% ethanol for 5 min and then treated with an alkaline-saturated salt solution. The stained sections were mounted in a medium compatible with organic solvents and observed under polarized light with a Zeiss microscope (Zeiss, Oberkochen, West Germany).
Morphology studies
Small blocks of amyloid deposits were fixed in 4% (w/v) paraformaldehyde (PFA) and 0.1% (v/v) glutaraldehyde (GA) in 0.1 mol/l phosphate-buffered saline (PBS), pH 7.4, for 2 h at 4°C, dehydrated in graded ethanols, and embedded in Lowicryl K4M (Chemische Werke Lowi, Wald-Kraiburg, Germany). Sections were transferred onto glass slides and stained with 0.5% methylene blue and 0.5% borax for 30 s at 90°C. The stained sections were then mounted in DPX medium and observed under a Polyvar 2 optical microscope (Reichert-Jung, Wien, Austria) under immersion oil.
For ultrastructural studies, after fixation with 4% PFA and 0.15 GA, small pieces of amyloid deposits were post-fixed in 2.5% GA (v/v) in PBS. The samples were then progressively treated in graded acetone-resin solutions, post-fixed with 0.1% osmium tetroxide, and embedded in Spurr (Agar Scientific Ltd, Stanstead, UK). Ultrathin sections were obtained using an Ultracut E system (Reichert-Jung) and counterstained with uranyl acetate and lead citrate before examination using a transmission electron microscope.
Immunogold methods
For the ultrastructural studies, blocks of approximately 1 mm3 of amyloid deposits were fixed and embedded in Lowicryl K4M. Ultrathin sections (6090 nm) were obtained using an Ultracut E system and mounted on formvar-coated and etched gold grids.
Before labelling, sections were rinsed twice with 0.1 mol/l glycinePBS for 5 min and incubated with 2% ovalbumin in PBS for 30 min to block unspecific antibodyantigen complexes. The grids with human ß2-microglobulin amyloidosis were then incubated with polyclonal anti-ß2-microglobulin (dilution 1:500; Nordic, Tilburg, The Netherlands) and diluted in 1% ovalbumin, and the grids containing murine AA amyloidosis were incubated with a polyclonal Ab against SAA (Dako, Glostrup, Denmark) and diluted in 1% ovalbumin for 2 h. After three 15-min rinses in glycinePBS, bound antibodies were visualized with 10 nm protein A gold (pAg) (kindly provided by Dr J. W. Slot, University of Utrecht, The Netherlands). The sections were finally rinsed with PBS and double distilled water prior to counterstaining with aqueous uranyl acetate and lead citrate. Controls were performed by omitting the primary antibody. Observations were carried out with Hitachi H-600 AB (Japan) and Philips EM-300 (The Netherlands) transmission electron microscopes.
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Results |
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Amyloid deposits in the liver were observed as fibril bundles located around the vessels, occupying the subendothelial space (Figure 1A). There was increased cellularity and thickening of the subendothelial space and the basal lamina, as well as the abnormal presence of collagen (Figure 1D
). Animals sacrificed after 4 weeks of amyloid induction (late stages of amyloidosis) presented amyloid deposits that occupied most of the analysed tissue, and showed a decrease in the cellular infiltrate. Liver tissue from a control animal is shown in Figure 1B
for comparison.
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ß2-Microglobulin amyloid deposits
Vascular involvement and membrane modifications were also studied in human ß2-microglobulin amyloidosis to determine whether these pathological findings observed in experimental amyloidosis also participate in dialysis-related amyloidosis. We observed findings that were very consistent among our patients. Amyloid deposits consisted of an intercellular amorphous material, including macrophage-like cells and blood vessels that supplied this abnormal tissue (Figure 2). They were clearly separated from the surrounding tissue by a loose connective matrix. The cells and amyloid fibrils were grouped in regions usually associated with a single blood vessel. These areas were separated from each other by a layer of connective tissue that included thin bundles of collagen fibres and some elastic fibres. These elastic fibres were characteristically associated with the highest ordered amyloid fibrils (Figure 3
).
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The vessels observed around amyloid deposits were mainly from precapillary sphincter regions, capillaries, and pericytic venulae. Arterioles and venulae irrigating the surrounding connective tissue were observed only occasionally (Figure 4). The capillaries and pericytic venulae showed an increased number of endothelial cells, the presence of buttons of cell proliferation (Figure 4B
), and enlargement of subendothelial spaces due to deposition of amyloid material (Figure 4A
). Macrophages and pericytes were associated with these vessel areas of amyloid deposits (Figure 4C
).
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Discussion |
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The location of amyloid fibrils in the subendothelial space has been classically observed in several types of amyloidoses, including hereditary cerebral haemorrhage amyloidosis [12] and Alzheimer's disease [13]. In ß2-microglobulin amyloidosis, there is no agreement about the vascular distribution of amyloid fibrils. Gal et al. [14], studying necropsy specimens from dialysis patients, found that amyloid material was preferentially observed in the wall of blood vessels of the osteoarticular system, gastrointestinal tract, heart, and lungs. In contrast, Noël et al. [15] failed to find a significant multiorgan involvement that included a vascular distribution of deposits in 23 patients with ß2-microglobulin amyloidosis. Our study showed the presence of amyloid deposits in the subendothelial space of blood vessels. However, we also found that amyloid deposits are not homogeneous in terms of amyloid fibril density, the presence of cells, and the proportion of blood vessels. These disparities may contribute to the differences reported by these authors.
The second aspect of vascular participation in the pathogenesis of amyloidosis is the carrier function. It is clear that for amyloid deposits to appear, the substrate and cofactors (normally proteins) as well as the cells need to reach the precise location where amyloidogenesis occurs. We previously reported the presence of albumin, transferrin, haemopexin, and other serum proteins in ß2-microglobulin amyloidosis [16] and Graeber et al. [17] identified pre-albumin, albumin, immunoglobulins, and amyloid P component, as well as complement factors in amyloid deposits from Alzheimer's disease patients. The presence of serum proteins in amyloid deposits suggests an increased permeability of blood vessels in these regions. Therefore, blood vessels not only provide a path to reach amyloid deposits, but the barrier function of the endothelium may also be altered, allowing serum proteins to reach the subendothelial and extravascular spaces and precipitate into amyloid fibrils.
Although cellular infiltrates have been less frequently studied than proteins in amyloidogenesis, certain details are well established. The density of these cells is relatively small (<200 cells/0.2 mm2) and >90% of them express the CD14 antigen, indicating that they are of macrophage origin [9]. Although macrophages have been thought to participate in the formation of amyloid fibrils [18,19], they may also contribute to a secondary process aimed at clearing the affected tissues of amyloid deposits by phagocytosis [11]. Regardless of whether macrophages are at the origin of amyloid fibril formation or whether they are a reactive phenomenon, the higher density of cells around blood vessels is in keeping with a vascular provenance of the cellular infiltrate.
In addition to providing a path for amyloid components to reach amyloid deposits and the altered barrier function of endothelium, blood vessels display characteristics suggesting an active participation in the disease. They have an increased number of cells, proliferation buttons, numerous transcytotic vesicles, and thickened or multi-layered basal lamina. These features, controlled by pericytes, are observed in angiogenesis and are associated with an increased vascular permeability [20].
Finally, our study showed that cellular membranes in close contact with amyloid fibrils were damaged. This was true for the nuclear envelope as well as for the cytosolic endomembranes, indicating a toxic effect of the amyloid fibrils that may result in cellular damage and death. We have recently shown that there is an impaired processing of ß2-microglobulin from amyloid fibrils occurring in the lysosomes of macrophages [11]. The unprocessed ß2-microglobulin is retained in the cell and can result in cell lysis. Our findings in experimental amyloidosis suggest a series of sequential events in amyloidogenesis. The animals analysed after 1 week of amyloid induction had spleen deposits in vascular areas showing a large proportion of cells. The animals having longer periods of amyloidosis presented with large clumps of amyloid fibrils and sparsely distributed blood vessels and cells. These findings are in agreement with a toxic effect of undigested amyloid proteins that would produce cellular death. As we have observed in ß2-microglobulin amyloidosis [11], the cells would then be replaced by amyloid fibrils which would become predominant in areas of amyloid deposits along with cellular debris that would form between amyloid fibrils.
In summary, the present findings suggest that blood vessels participate in amyloidogenesis through a variety of mechanisms: blood vessels are sites of amyloid fibril deposition, they facilitate the access of the serum proteins and blood cells into amyloid deposits, and they react with cell proliferation and angiogenesis factors. Our sequential analysis of experimental murine amyloidosis supports a toxic effect of amyloid fibrils that produce cell death followed by immersion of cellular debris in accumulated amyloid fibrils.
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Acknowledgments |
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Notes |
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References |
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