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*
Division of Nephrology, Université
Catholique de Louvain Medical School, Brussels, Belgium
Department of Pathology, Université
Catholique de Louvain Medical School, Brussels, Belgium
Division of Cell Biology, Commissariat à
l'Energie Atomique, Saclay, France
§
Institute of Medical Science and Department of Internal Medicine, Tokai
University School of Medicine, Kanagawa, Japan
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Division of Nephrology, Centre Hospitalier de Luxembourg,
Luxembourg.
Correspondence to Dr. Olivier Devuyst, Division of Nephrology, Université Catholique de Louvain, 10 Avenue Hippocrate, B-1200 Brussels, Belgium. Phone: +32 2 764 1855; Fax: +32 2 764 2836; E-mail: devuyst{at}nefr.ucl.ac.be
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Nitric oxide (NO) plays a key signaling role in countless biologic processes, including control of vascular tone and permeability (11,12), as well as angiogenesis, via an interaction with vascular endothelial growth factor (VEGF) (13,14). Nitric oxide is synthesized from L-arginine by a family of three NO synthase (NOS) isozymes, named from the tissue in which they were initially cloned: neuronal NOS (nNOS), inducible NOS (iNOS; cloned from macrophages), and endothelial NOS (eNOS) (15). Specific NOS isoforms are expressed in human and rat peritoneum (16,17), and detection of nitrite and nitrate in the dialysate attests to the occurrence of NOS activity in the peritoneum (18,19). Several lines of evidence suggest that NO is involved in the regulation of peritoneal transport during PD, with or without peritonitis. Addition of the NO donor nitroprusside to the dialysate increases both effective peritoneal surface area and intrinsic permeability of the membrane in animal models (20) or stable PD patients (21). In acute peritonitis, decreased UF is associated with a major increase in peritoneal NOS activity, due to upregulation of both eNOS and iNOS (17). However, it is still unknown whether the production of NO and the expression of the various NOS isoforms in the peritoneum are modified in long-term PD.
We have thus tested the hypothesis that modifications of activity and/or expression of NOS isozymes play a role in the increased effective peritoneal surface area observed in long-term PD patients. Using the L-citrulline assay, we determined the specific activities of NOS isoforms in a series of peritoneal biopsies obtained from control subjects and uremic patients treated with PD. These data were correlated with: (1) morphometric quantification of vascular density and endothelial area; (2) immunoblotting studies for NOS isoforms and VEGF; and (3) immunostaining for eNOS, VEGF, and the advanced glycation end product (AGE) pentosidine.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Tissue Processing
Peritoneal biopsies were obtained at surgery after careful dissection with
scissors, with minimal mechanical contact and no electrocoagulation. This
technique allows for the preservation of an excellent integrity of the tissue
(16). Samples were folded
inside-out to avoid abrasion of the mesothelium and washed in ice-cold
phosphate-buffered saline (PBS). If size was sufficient, a small part of the
samples was fixed in 4% paraformaldehyde in PBS (pH 7.4) and further embedded
in paraffin (16). The major
part of the samples was snapfrozen in liquid nitrogen to perform protein
extraction as described previously
(16,
17). Briefly, samples were
grounded in liquid nitrogen and suspended in 2 ml/g ice-cold homogenization
buffer (50 mM Tris, pH 7.4, containing 0.1 mM
ethyleneglycol-bis(ß-aminoethyl ether)-N,N'-tetra-acetic
acid [EGTA], 0.1 mM ethylenediaminetetra-acetic acid [EDTA], 2 mM
ß-mercaptoethanol, 5 µM leupeptin, and 4 µM pepstatin). The
suspension was further homogenized with an Ultra-Turrax® (Labortechnik,
Staufen, Germany) and then briefly sonicated (Branson Sonifier B12, Danbury,
CT). The resulting homogenate was centrifuged at 6000 x g
(Sigma 113 Centrifuge, Osterode am Harz, Germany) for 10 min at 4°C. After
determination of protein concentration with the Bradford method (Bio-Rad), the
post-nuclear supernatant (total protein extract) was kept at -80°C.
Measurement of NOS Activity
NOS activity was determined by the conversion of L-[3H]-arginine
to L-[3H]-citrulline as described previously
(17,
23). Briefly, 25 µl of
tissue extract (approximately 250 to 400 µg of total protein) containing 20
mM 3-[(3-cholamidopropyl)dimethylamino]-1-propane sulfonate was added to 200
µl of Tris buffer (50 mM, pH 7.4) containing 10 mM dithiothreitol, 10 µM
tetrahydrobiopterin, 10 µg/ml calmodulin, 1 mM NADPH, 4 µM flavin
adenine dinucleotide, 4 µM flavin mononucleotide, 2 µM L-arginine, and
10-3 mCi/ml L-[3H]-arginine. Assays were performed for
30 min at 37°C and terminated with 2 ml of ice-cold stop buffer (20 mM
CH3COONa, pH 5.5, containing 2 mM EDTA, 0.2 mM EGTA, and 1 mM
L-citrulline). L-[3H]-Citrulline was separated from the incubation
mixture by cation exchange chromatography, using Dowex AG 50W-X8 resin
(Bio-Rad) and, after elution with water, quantified by liquid scintillation
counting (SL3000; Intertechnique, Plaisir, France). The NOS activity (in pmol
citrulline/mg protein per min) was determined from counts obtained with and
without 1 mM NG-monomethyl-L-arginine, a specific
inhibitor of NOS. When a sufficient amount of sample was available, assays
were performed with Ca2+ (1 mM CaCl2) or without
Ca2+ (0 mM CaCl2, 2 mM EGTA, and 2 mM EDTA) to measure
total versus Ca2+-independent NOS activities,
respectively. The Ca2+-dependent NOS activity was obtained by
subtracting Ca2+-independent NOS activity from total NOS activity.
Results were normalized for protein content. Determination of NOS enzymatic
activity as a function of protein concentration, temperature, and time
verified that the assay was made in the linear part of the curves
(23). The determinations were
performed in duplicate.
Antibodies
NOS isoforms were detected with mouse monoclonal antibodies raised against
human eNOS and nNOS, and mouse iNOS (Transduction Laboratories, Lexington,
KY). Specificity of these antibodies against NOS isoforms has been
demonstrated (17). Other
antibodies included a purified rabbit anti-human factor VIII IgG (Dakopatts,
Glostrup, Denmark), a rabbit polyclonal antiserum against nitrotyrosine
(Upstate Biotechnology, Lake Placid, NY)
(24), a monoclonal antibody
against human VEGF (Santa Cruz Biotechnology, Santa Cruz, CA)
(25), and an affinity-purified
rabbit antibody against pentosidine
(26).
Other Reagents and Supplies
Peroxidase-labeled goat anti-rabbit IgG and goat anti-mouse IgG were from
Dako (Glostrup, Denmark); rabbit and mouse IgG avidinbiotin peroxidase complex
(ABC) kits were from Vector Laboratories (Burlingame, CA). Electrophoresis
reagents were from Bio-Rad (Melville, NY), and enhanced chemiluminescence was
from Amersham (Arlington Heights, IL). L-[3H]-Arginine was from
Amersham (Buckinghamshire, United Kingdom) and liquid scintillation reagent
was from Lumac (Groningen, The Netherlands). Other reagents and supplies were
from Sigma Chemical Co. (St. Louis, MO), J. T. Baker (Phillipsburg, NJ),
National Diagnostics (Atlanta, GA), Boehringer (Mannheim, Germany),
Polysciences (Warrington, PA), and Pierce (Rockford, IL).
Immunoblot Analyses
Sodium dodecyl sulfate-polyacrylamide gel electrophoresis and
immunoblotting were performed as described earlier
(16,
17). The extracts were
solubilized by heating (95°C for 2 min) in sample buffer (1.5% sodium
dodecyl sulfate, 10 mM Tris-HCl, pH 6.8, 0.6% dithiothreitol, and 6% [vol/vol]
glycerol). Proteins (40 µg/lane) were separated by electrophoresis through
0.1 x 9 x 6 cm 7.5% or 12% acrylamide slabs and transferred to
nitrocellulose. After Ponceau Red (Sigma) staining to check transfer
efficiency, destained membranes were blocked for 30 min at room temperature in
blocking buffer (50 mM sodium phosphate buffer, 150 mM NaCl, 0.05% Tween 20,
pH 7.4) comprising 5% nonfat dry milk, followed by incubation with the primary
antibody (diluted in blocking buffer with 2% bovine serum albumin) at 4°C
for 16 to 18 h. The membranes were then washed, incubated for 1 h at room
temperature with the appropriate peroxidase-labeled antibody (1:5000
dilution), washed again, and visualized with enhanced chemiluminescence
(Amersham). The specificity of the immunoreaction was determined by comparison
with the signal observed with positive controls and incubation with nonimmune
rabbit or mouse IgG (Vector). Densitometry analysis was performed with a
StudioStar Scanner (Agfa-Gevaert, Mortsel, Belgium), using the NIH-Image V1-57
software. Optical densities (given in arbitrary densitometry units) were
obtained in duplicate from two different gels.
Immunohistochemistry
Immunoperoxidase staining on human peritoneum sections was performed as
described previously (16,
17). Six-micrometer sections
were cut from paraffin blocks, dewaxed, and rehydrated in graded ethanols.
After inhibition of endogenous peroxidase by incubation in 0.3%
H2O2 for 30 min, the slides were blocked with 10% normal
goat or horse serum in PBS for 20 min at room temperature. All subsequent
antibody incubations were carried out for 45 min at room temperature in a
humidified chamber. Sections were incubated with the primary antibody diluted
in PBS containing 2% bovine serum albumin, washed 3 x 5 min, incubated
with biotinylated goat anti-rabbit or horse anti-mouse IgG (Vector), washed 3
x 5 min, and then incubated for 45 min with the avidin-biotin peroxidase
complex (Vector). After washing for 3 x 5 min, antibody localizations
were visualized using aminoethylcarbazole. For pentosidine staining,
rehydrated sections were incubated in a buffer (0.05 M Tris-HCl, pH 7.2, 0.1 M
NaCl) containing 0.5 mg/ml Pronase (Dako) for 15 min at room temperature,
washed in PBS, blocked in 4% milk for 2 h, and subsequently incubated with the
anti-pentosidine rabbit IgG
(26) overnight at 4°C.
After washing, sections were incubated with a peroxidase-conjugated antibody,
and detection was performed with 3,3'-diaminobenzidine containing 0.003%
H2O2. Sections were viewed under a Leica DMR coupled to
a Leica MPS60 photomicrographic system (Leica, Heerbrugg, Switzerland). The
specificity of the immunolabeling was confirmed by: (1) incubation
without primary or secondary antibody; and (2) incubation with
nonimmune rabbit or mouse IgG (Vector).
Quantification of Vascular Immunoreactivity for Factor VIII
Quantification of factor VIII reactivity was performed by image analysis
(27). After staining for
factor VIII, sections of human peritoneum were systematically examined through
a Zeiss microscope using a KS-400 image analysis system (Kontron, Munich,
Germany) coupled to a DAGE-MTI charge-coupled device 72 camera (Michigan City,
IN). The stained peritoneum areas were digitized to allow automated detection
of stained structures and selection of the objects corresponding to vessel
walls. Analyses included determination of: (1) the density of stained
vascular structures (in n vessels/field); (2) the mean
vessel area, defined as the mean area of the lumen of the blood vessels
stained for factor VIII (in µm2); and (3) the
cumulative vascular area and the cumulative endothelial area (both in % of the
fields examined). All slides were coded and analyzed on a single-blind basis
by the same operator.
Quantification of Pentosidine Levels
Pentosidine content in human peritoneum was assessed by HPLC assay as
described previously (28).
Briefly, the peritoneal sample was lyophilized and then hydrolyzed in 50 µl
of 6N HCl for 16 h at 110°C under nitrogen, followed by neutralization
with 50 µl of 5N NaOH and 100 µl of 0.5 M phosphate buffer (pH 7.4),
then filtered through a 0.5-µm filter and diluted with PBS. A sample
(corresponding to approximately 20 µg of protein) was injected into an HPLC
system and fractionated on a C18 reversed-phase column (Waters, Tokyo, Japan).
The effluent was monitored at an excitation-emission wave-length of 335/385 nm
using a fluorescence detector (RF-10A, Shimadzu, Japan). Synthetic pentosidine
(29) was used to obtain a
standard curve. The identity of the substance in the specimens, detected at
the same retention time as authentic pentosidine, was confirmed as pentosidine
by fast-atom bombardment-mass spectrometry (379.4 Daltons). Limits of
detection were 0.1 pmol of pentosidine per milligram of protein.
Statistical Analyses
Data are presented as mean ± SEM. Statistical significance of the
differences between groups was assessed using t test (two means) or
ANOVA (three or more means), as appropriate.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The L-citrulline assay was performed in parallel with and without Ca2+ to determine which NOS isoform (Ca2+-dependent and/or Ca2+-independent) was involved. This analysis was performed in five control subjects and in five short-term and six long-term PD patients in whom sufficient peritoneal extract was available. As shown in Figure 2, the increase in total NOS activity observed in long-term PD patients was solely due to an increase in Ca2+-dependent NOS activity, which averaged 0.10 ± 0.02 pmol citrulline/mg protein per min in long-term PD patients, compared to 0.015 ± 0.01 in control subjects and 0.03 ± 0.01 in short-term PD patients (Figure 2A). In contrast, the Ca2+-independent NOS activity (Figure B) was similar in long-term PD patients, short-term PD patients, and control subjects (0.03 ± 0.01, 0.02 ± 0.01, and 0.015 ± 0.01 pmol citrulline/mg protein per min, respectively).
Expression of NOS Isoforms in Human Peritoneum: Immunoblot
Analysis
Immunoblot analysis was performed to identify the isoform, eNOS or nNOS,
implicated in the increased Ca2+-dependent NOS activity observed in
long-term PD patients. Extracts from bovine aortic endothelial cells and rat
pituitary gland were used as positive controls for eNOS (140 kD) and nNOS (155
kD), respectively. As demonstrated on the representative immunoblot shown in
Figure 3A, a weak band
corresponding to eNOS was detected with a variable intensity in control
subjects and short-term PD patients; that band was significantly upregulated
in peritoneum extracts from long-term PD patients (lanes 3 to 6). Optical
densitometry analyses confirmed a significant, approximately threefold
increase of eNOS expression in the peritoneum of long-term PD patients
compared with control subjects and short-term PD patients
(Figure 3B). In contrast with
eNOS, no specific signal for nNOS could be detected in the peritoneal samples
examined, even with longer film exposure
(Figure 3A). No signal was
detected when immunoblots were probed with nonimmune, control mouse IgG, or
with a monoclonal anti-iNOS at the same dilution (data not shown).
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Immunolocalization of eNOS and Nitrotyrosine in the Peritoneum of
Control Subjects and PD Patients
A relatively faint immunostaining for eNOS was detected in the parietal
peritoneum from both control subjects and long-term PD patients
(Figure 4, A through D). As
described previously (16), the
signal for eNOS was located in the endothelium lining peritoneal blood
vessels. Compared with control subjects
(Figure 4, A and C), the
staining was more intense in the peritoneum of long-term PD patients
(Figure 4, B and D). No
staining was detected in sections incubated with nonimmune, control mouse IgG
at the same dilution (Figure
4E). No specific staining for nNOS could be observed in the
peritoneum of both control subjects and PD patients
(Figure 4, F and G). The
immunoreactivity for nitrotyrosine was tested to investigate whether the
increased NOS activity in long-term PD patients was associated with
peroxynitrite formation and nitration of tyrosine residues. The peritoneal
staining for nitrotyrosine was consistently increased in long-term PD
patients, particularly along peritoneal capillaries
(Figure 4, compare H and I). No
specific staining was observed when the same sections were incubated with
control rabbit IgG (data not shown).
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Quantification of Vascular Immunoreactivity for Factor VIII
Quantitative studies investigated whether the increased expression of eNOS
in the peritoneum of long-term PD patients correlated with a proliferation of
blood vessels within the peritoneum. Immunostaining for factor VIII
(31), performed in
representative peritoneal sections from control subjects and long-term PD
patients, showed a net increase in the density of stained structures in the
latter (Figure 5).
Quantification of factor VIII immunoreactivity
(Table 1) showed a significant,
2.5-fold increase in the density of stained vessels in long-term PD patients
compared with control subjects. The mean area of the stained vessels (the area
included in the labeling, i.e., the lumen plus the subendothelial
area) was also slightly increased in PD patients. As a result, there was a
significant increase of both the cumulative endothelial area and the
cumulative vascular area in long-term PD patients. This morphometry analysis
has been validated on serial sections of the peritoneum, with a mean
interassay variability of 16 ± 8% (density parameter).
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Expression of VEGF and Pentosidine
Expression studies for VEGF were conducted to address the putative role of
that growth factor in the vascular proliferation observed in the peritoneum of
long-term PD patients. Immunoblot analysis
(Figure 6) showed a strong
signal for VEGF in most short-term and long-term PD patients, whereas no
signal was detected in control subjects. Immunostaining for VEGF in the
peritoneum confirmed a marked increase of the labeling, mostly along capillary
endothelium, in long-term PD patients versus control subjects
(Figure 7, A and B). A signal
for VEGF was also detected in the mesothelium
(Figure 7, A and B). No
staining was observed when sections were incubated with nonimmune mouse IgG
(Figure 7C).
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To substantiate a putative link between carbonyl stress, AGE deposition in the peritoneum, and VEGF secretion by peritoneal cells (32, 33), we determined the level of pentosidine in peritoneal biopsies obtained in control subjects and PD patients. The pentosidine level was below the detection limit (<0.1 pmol/mg protein) for all control subjects tested (n = 5), while a mean level of 11 ± 8 pmol/mg protein (range, 0.2 to 36.1 pmol/mg protein) was detected in the peritoneum of tested long-term PD patients (n = 4). This difference in pentosidine levels was reflected by immunostaining for pentosidine in representative sections of the peritoneum (Figure 8). A weak but significant staining for pentosidine was indeed detected in the peritoneum of long-term PD patients, whereas it was absent in control subjects (Figure 8, A and B). Staining for pentosidine in the peritoneum of PD patients was located both in the endothelium and mesothelium. The use of serial sections from a long-term PD patient demonstrated that staining for pentosidine (Figure 8C) colocalized with staining for VEGF (Figure 8D) in the endothelium.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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As pointed out in the introduction, the most frequent transport abnormality developing during long-term PD is UF failure (2, 33), a complication that eventually leads to the transfer of the patient to another modality of renal replacement therapy (33). The pathophysiology and the molecular mechanisms involved in the UF failure associated with long-term PD are unclear. Recurrent episodes of peritonitis, with associate inflammatory changes, may contribute to peritoneal damage over time (10), but are not a prerequisite for development of UF failure (9, 33). Functional studies have demonstrated that the loss of UF is associated with an increased transport of small solutes such as urea and glucose, and an attendant enhanced dissipation of the osmotic gradient across the peritoneal membrane (5, 6, 34). Modeling studies have suggested that the increased solute transport is mediated by an augmented peritoneal surface area (22). Because the endothelium lining peritoneal blood vessels represents the most important barrier to small solute transport (22), the expanded peritoneal surface must correspond to an increased vascular area (33). Indeed, preliminary reports have documented an increased vascular density in the peritoneum of long-term PD patients (35).
The present results provide a morphologic and molecular basis for this hypothesis. A significant fivefold increase in NOS activity is measured in the peritoneum of long-term PD patients compared with control subjects. Furthermore, in uremic patients, NOS activity is positively correlated with the duration of PD (Figure 1). The results of the global L-citrulline assay in the peritoneum are entirely due to a Ca2+-dependent NOS isozyme (Figure 2), which has been further characterized as eNOS by immunoblotting and immunostaining (Figures 3 and 4). The increased NOS activity in the peritoneum is highly specific, and these results differ from those obtained during acute peritonitis, a condition associated with upregulation of both eNOS and iNOS (17). No clinical and biologic signs of acute peritonitis were present in our patients, and absence of peritonitis is further attested by the lack of variation in Ca2+-independent NOS activity and the absence of signal for iNOS in the samples tested.
The combined increase in NOS activity and eNOS expression suggests that enhanced NO production might contribute to the progressive loss of peritoneal UF, via a vasodilation-induced rise in peritoneal vascular area and a subsequent dissipation of the osmotic gradient. Several lines of evidence support that hypothesis. In physiologic conditions, eNOS is the prevailing form of NOS in the vascular system, and it is indeed significantly expressed in the peritoneum vasculature (16). Our studies in human and rat peritoneum have shown that eNOS is markedly upregulated in peritoneal inflammation (16) or acute peritonitis (17). In the latter condition, increased NOS activity is negatively correlated with UF capacity (17, 36). Furthermore, the NO donor nitroprusside increases effective peritoneal surface area and intrinsic permeability (20, 21), whereas NOS inhibitors such as NG-nitro-L-arginine methyl ester decrease these parameters (37).
It must be emphasized that another potentially detrimental consequence of increased NO levels within the peritoneum might arise from biochemical modifications, including covalent modifications of proteins. As a radical, NO reacts with oxygen, superoxide anions, and transition metals, leading to S-nitrosylation of critical cysteine or tyrosine residues (38). These modifications are well illustrated by the increased staining for nitrotyrosine that characterizes the peritoneum of long-term PD patients (Figure 4). It is tempting to speculate that these modifications might play a role in the structural changes of the peritoneum (such as interstitial fibrosis or nitrosylation of endothelial proteins) that occur with time on PD (33, 39).
Several factors might explain the increased expression of eNOS in long-term
PD patients. Specific immunohistochemistry shows that eNOS is located in the
endothelium lining peritoneal blood vessels. The vascular density is clearly
augmented in long-term PD patients, as reflected by quantitative morphometry
after staining for factor VIII (Figure
5 and Table 1).
These results thus confirm a preliminary study
(35) and suggest that eNOS
upregulation might reflect vascular proliferation and increased endothelial
surface area. Alternatively, it is conceivable that eNOS expression in the
peritoneum might be modulated by changes in immune defense systems induced by
long-term PD, e.g., local cytokine generation
(40). The observations that
eNOS is upregulated in the inflammatory peritoneum
(16) or upon virus-induced
activation of interferon-
and tumor necrosis factor-
(41) support the latter
hypothesis.
The relationship between eNOS regulation and angiogenesis deserves further consideration. Of course, as mentioned earlier, increased eNOS may be a simple consequence of angiogenesis and increased endothelial area. However, increased NO levels might also be at the origin of angiogenesis within the peritoneum. Recent studies have shown that: (1) eNOS expression and angiogenesis are correlated during embryogenesis (42); (2) NO can induce angiogenesis in vivo (43); and (3) NO is necessary for the biologic activity of VEGF (13, 14). We now document that peritoneal angiogenesis is associated with an augmented expression of VEGF—a hitherto unreported characteristic of long-term PD. Immunostaining for VEGF showed a marked increase of the labeling, mostly along capillary endothelium, compared with control samples. These morphologic findings are confirmed by immunoblot analysis of peritoneal samples. Thus, enhanced VEGF expression together with an augmented NO release might play a determining role in the neoangiogenesis and its subsequent increase in peritoneal vascular area.
In an effort to identify the cause of the enhanced VEGF expression, we investigated the relationship between the latter phenomenon and the well-known AGE accumulation in the peritoneum during long-term PD (44,45,46). Through an interaction with their receptors, AGE induce the release of cytokines and growth factors (47). Furthermore, glucose degradation products such as methylglyoxal promote the transcription of VEGF by endothelial and mesothelial cells (32). We concentrated on one AGE structure, pentosidine, and document significantly higher levels of pentosidine in peritoneal extracts obtained from long-term PD patients than in control subjects. Furthermore, immunohistochemistry reveals a marked increase in pentosidine at both the endothelial and mesothelial levels of the peritoneum in long-term PD patients, as well as a colocalization with VEGF at the endothelial level (Figure 8). It is thus tempting to speculate that progressive AGE deposits on endothelial (and maybe mesothelial) cells within the peritoneum might promote liberation of VEGF, the latter stimulating angiogenesis in association with eNOS (13, 14, 48).
In conclusion, our data provide a morphologic (vascular proliferation and increased endothelial area) and molecular (eNOS upregulation and increased NOS activity) basis for explaining the increased peritoneal transport and surface area associated with long-term PD. Additional studies should focus on the implication of AGE and VEGF in that detrimental process.
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Acknowledgments |
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These studies were supported in part by the Fonds National de la Recherche Scientifique (crédit 9.4540.96), the Fonds de la Recherche Scientifique Médicale (convention 3.4566.97), and grants from Baxter. We thank Ch. van Ypersele, J.-L. Balligand, N. Lameire, and J.-M. Pochet for providing invaluable suggestions and critiques, as well as J.-P. Squifflet, J. Malaise, and M. Mourad for help in providing peritoneal biopsies. In particular, we thank the many patients and nurses without whom this study would not have been possible. The expert technical assistance of M. de Rudder, S. Ruttens, and L. Wenderickx is highly appreciated.
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Footnotes |
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American Society of Nephrology
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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, n = 5) and
short-term PD (
, n = 5) and long-term PD ([UNK], n =
6) patients. The Ca2+-dependent NOS activity is significantly
increased in long-term PD patients versus control subjects and
short-term PD patients (ANOVA: P = 0.001; *P <
0.003, long-term PD versus control subjects; #P =
0.01, long-term PD versus short-term PD), whereas
Ca2+-independent NOS is similar in the three groups. There are no
significant differences between short-term PD patients and control
subjects.
, n = 4
individual samples; [UNK], n = 2 pooled samples); and short-term
(
