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Am J Physiol Heart Circ Physiol 292: H285-H294, 2007. First published September 8, 2006; doi:10.1152/ajpheart.00560.2006 Free Article
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Chronic NOS inhibition actuates endothelial-mesenchymal transformation

Edmond O’Riordan, Natalia Mendelev, Susann Patschan, Daniel Patschan, Jonathan Eskander, Leona Cohen-Gould, Praveen Chander, and Michael S. Goligorsky

Departments of Medicine, Pharmacology, and Pathology, and Renal Research Institute, New York Medical College, Valhalla, New York; and Cornell University, Weill Medical College, New York, New York

Submitted 31 May 2006 ; accepted in final form 29 August 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Chronic kidney diseases are accompanied by the accumulation of substances like asymmetric dimethylarginine, phenylacetic acid, homocysteine, and advanced glycation end products, known to either inhibit endothelial nitric oxide synthase (eNOS) or uncouple it, consequently limiting the amount of available nitric oxide (NO). Reduced bioavailability of NO induces endothelial dysfunction. An early loss of peritubular capillaries in tubulointerstitial fibrotic areas and injury to endothelial cells have been linked to progressive renal disease. Screening endothelial genes in cells treated with NOS inhibitors showed upregulation of collagen XVIII, a precursor of a potent antiangiogenic substance, endostatin. This finding was confirmed at the level of mRNA and protein expression. Tie-2 promoter-driven green fluorescent protein mice treated with nonhypertensinogenic doses of a NOS inhibitor exhibited upregulation of collagen XVIII/endostatin and rarefaction of capillary profiles. This was accompanied by the increased expression of transforming growth factor-beta and connective tissue growth factor in the kidney. Occasional endothelial cells expressed both the marker of endothelial lineage (green fluorescent protein) and mesenchymal marker ({alpha}-smooth muscle actin or calponin). In vitro studies of endothelial cells treated with asymmetric dimethylarginine showed decreased expression of eNOS and Flk-1 and enhanced expression of calponin and fibronectin, additional markers of smooth muscle and mesenchymal cells. These cells overexpressed transforming growth factor-beta and connective tissue growth factor, as well as endostatin. In conclusion, data presented here 1) ascribe to NO deficiency in endothelial cells the function of a profibrotic stimulus associated with the expression of an antiangiogenic fragment of collagen XVIII (endostatin) and 2) provide evidence of endothelial-mesenchymal transdifferentiation in the course of inhibition of NOS by a pathophysiologically important antagonist, asymmetric dimethylarginine. Both mechanisms may account for microvascular rarefaction.

endothelial dysfunction; endostatin; transforming growth factor-beta; connective tissue growth factor

GLOMERULOSCLEROSIS AND TUBULOINTERSTITIAL scarring are the main processes underlying progression of chronic renal diseases, regardless of their etiology (reviewed in Refs. 5, 12, 13, 17, 33, 44). There is substantial evidence attributing these pathological processes to such mechanistic factors as hypertension and ANG II, proteinuria, oxidative stress, advanced glycation end products, tubular obstruction and increased tubular pressure, various growth factors, especially transforming growth factor (TGF)-beta and PDGF, and elevated tissue plasminogen activator and plasminogen activator inhibitor 1. Some of these mechanistic factors have been implicated in epithelial-mesenchymal transition with accumulation of myofibroblasts and interstitial-type collagens in nephrosclerosis. An early loss of peritubular capillaries in tubulointerstitial fibrotic areas (4) and injury to endothelial cells (34) have been proposed as mechanisms of progressive renal disease. Recent studies provide a solid experimental basis for the concept of microvascular injury and chronic ischemia as a prerequisite for progression of fibrosis in etiologically diverse renal diseases (27–30, 46, 47). Although it has been firmly established that nephrosclerosis is accompanied by microvascular rarefaction, its pathogenesis remains obscure. We hypothesized that endothelial-mesenchymal transformation, analogous to the epithelial-mesenchymal transition, may contribute to microvascular rarefaction and ensuing chronic renal ischemia.

Chronic kidney diseases are accompanied by the accumulation of substances like asymmetric dimethylarginine (ADMA), phenylacetic acid, homocysteine, and advanced glycation end products, known to either inhibit endothelial nitric oxide synthase (eNOS) or uncouple it, consequently limiting the amount of available nitric oxide (NO) (6, 16, 25, 32, 38, 39, 55, 58). Reduced bioavailability of NO is a cornerstone of the developing endothelial dysfunction. It is instructive that therapeutic strategies proposed and used to halt the progression of nephrosclerosis, such as angiotensin-converting enzyme inhibition, activation of bradykinin B2 receptor, L-arginine supplementation, and combinations of lisinopril and L-arginine (50, 53, 59), are in fact agonists of eNOS. Indeed, eNOS function appears to be a good predictor of individual susceptibility to renal damage in subtotally nephrectomized rats; it also protects Wistar Furth rats from chronic renal injury (14, 21). Furthermore, previous observations in experimental animals receiving NOS inhibitors showed the development of nephrosclerosis and chronic renal insufficiency (3, 52, 57). This was the reasoning behind the present investigation into the primary role of endothelial dysfunction induced by eNOS inhibition as a potential mechanism for vascular rarefaction and nephrosclerosis.

In preliminary experiments using cDNA microarray, we screened the cardiovascular-relevant genes of cultured human umbilical vein endothelial cells (HUVEC) subjected to a NOS inhibitor. The results of this screening revealed upregulation of collagenase XI (2-fold), collagenase IV ({alpha}3-chain) (1.9-fold), collagenase XVIII (1.9-fold), matrix metalloproteinase-1 (1.5-fold), {alpha}2-integrin (2-fold), and fibrinogen (2.5-fold) and downregulation of tissue inhibitor of metalloproteinase-4 gene (2.4-fold), thus alluding to the potential participation of the endothelium in fibrotic processes. Further preliminary analyses of these findings by RT-PCR and immunocytochemistry of cultured cells, as well as studies in an in vivo model, demonstrated that it was predominantly collagen XVIII, a precursor of the endogenous potent anti-angiogenic substance endostatin (proteolytic COOH-terminal fragment of collagen XVIII) (48), that was consistently upregulated in all these screens. Hence, endostatin/collagen XVIII was chosen as the most promising target for the present investigation. The approach that we have developed was two pronged: 1) to validate the microarray findings using an in vitro system and 2) to extend the observations made in vitro to an in vivo model. Therefore, we examined the in vitro development of an altered phenotype in HUVEC treated with the NOS inhibitors NG-nitro-L-arginine methyl ester (L-NAME) or ADMA. In vivo experiments that immunohistochemically assessed endostatin/collagen XVIII expression in Tie-2 promoter-driven green fluorescent protein (Tie-2/GFP) transgenic mice treated with L-NAME [as well as in eNOS and neuronal NOS knockout mice] were also performed to further examine this hypothesis. A low dose of L-NAME was used to ensure that the blood pressure remained in the normal range, thus negating the known profibrotic effect of hypertension on the kidney. Accumulation of endostatin/collagen XVIII was demonstrated in cultured endothelial cells and nonhypertensive mice receiving L-NAME. This was associated with upregulation of TGF-beta and connective tissue growth factor (CTGF) and transformation of endothelial cells to mesenchymal phenotype.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Materials. The following antibodies were used: anti-collagen XVIII (sc-16651), anti-CTGF L-20 (sc-14939), anti-TGF-beta1/2/3 (sc-7892), anti-fibronectin (sc-9068), anti-Tie-2 (sc-9026), anti-CD31 (sc-1506), and anti-Flk-1 (sc-505), all from Santa Cruz Biotechnology. Anti-{alpha}-smooth muscle actin ({alpha}-SMA) 1A4 (M0851) antibodies were obtained from Dako Cytomation. Antibodies to calponin (c-2687) and endostatin (E-3779) were from Sigma (St. Louis, MO). Monoclonal antibodies to beta-actin were obtained from Sigma. L-NAME, NG-nitro-D-arginine methyl ester (D-NAME), and ADMA were obtained from Alexis Biochemicals (San Diego, CA). Endostatin (human recombinant, expressed in Pichiia pastoris) was purchased from Sigma, and human recombinant TGF-beta2 was from PeproTech (Rocky Hill, NJ).

Cell culture. HUVEC were cultured in endothelial cell basal medium 2 (Clontech) and used for experiments at passages 4–6. Cells were treated with L-NAME (150 µM) or equal concentrations of D-NAME. ADMA was used at a concentration of 5–15 µM and was added daily for the duration of 48–72 h in culture. Total RNA was isolated by TRIzol reagent (Invitrogen) after 24 h of treatment with L-NAME. Protein extraction was performed after 48–72 h of eNOS inhibition, using radioimmunoprecipitation buffer.

Quantitative RT-PCR. RT-PCR was performed by a GeneAmp 2700 PCR system (Applied Biosystems). RT-PCR protocol was as follows: RT step at 50°C for 30 min and 94°C for 2 min; and PCR thermal cycle at 94°C for 10 s, 65°C for 60 s, and 72°C for 120 s (30 cycles) with a final elongation for 7 min. Primers used were as follows: for collagen XVIII, AGCTGGGTCAGGCAGGGTGCAG (sense) and ATGGCCTTGTGCTGAGTGTGGCC (antisense). Ribosomal 18S primers were obtained from Ambion. Avian myeloblastosis virus reverse transcriptase Titan One Tube (Roche) and Superscript II (Invitrogen) were used according to manufacturer’s instructions.

Western blotting. Western blotting was performed with an XCELL-II blot module and 4–20% Tris-glycine gels (Novex). Cells were lysed in radioimmunoprecipitation buffer, and 40 µg of total protein were separated by electrophoresis on SDS-PAGE, followed by the transfer to Immobilon-P membrane. Membranes were incubated with antibodies to endothelial and mesenchymal markers, as detailed in RESULTS. Antibody binding was visualized with SuperSignal West Pico chemiluminescent substrate (Pierce).

Animal models. The animal study protocol was approved by our Institutional Animal Care and Use Committee. FVB and Tie-2/GFP mice on FVB background were obtained from Jackson Laboratories (Bar Harbor, ME), and neuronal NOS- and eNOS-deficient mice were kindly provided by T. Hintze (New York Medical College). Tie-2/GFP mice express GFP driven by the endothelial-specific and selective promoter for Tie-2 receptor, resulting in specific fluorescence of predominantly endothelial cells, as previously described (40). All animals had free access to water and food throughout the study. Animals were separately caged with light-dark cycle of 12:12 h and received NOS inhibitor or an inactive analog with the drinking water. Mice were randomly divided into five treatment groups: group A (control), group B (0.1 g/lL-NAME in drinking water), group C (0.4 g/lL-NAME in drinking water), group D (0.1 g/lD-NAME in drinking water), and group E (0.4 g/lD-NAME in drinking water). L-NAME doses of 0.4 mg/ml are presented as the maximal nonhypertensive dose, with equivalent doses of D-NAME.

Blood pressure measurements. Tail-cuff blood pressure was measured in conscious mice according to the previously described protocol (51). A fixed warming period of 10 min was used for each animal. All animals had 2 days of blood pressure measurement "training," after which blood pressure measurements were performed for 2 consecutive days (baseline) Blood pressure was recorded 10 times in each animal at every session, and 8 of these readings were averaged, with the highest and lowest readings being discarded. Blood pressure recording was then repeated three times during the 7- to 8-wk treatment period and before death.

Histochemical and immunofluorescence analyses. Tissue samples for frozen sections were fixed in periodate-lysine-paraformaldehyde buffer for 24 h and then immersed in sucrose. The tissue was then embedded in OCT (Tissue-Tek, Torrance, CA), and 5-µm-thick sections were prepared by Leica CM1850 microtome. The number of capillary profiles was estimated on frozen sections by counting the number of GFP-fluorescent endothelial cells per 100 tubules or glomeruli in a high-power field (x400).

Paraffin blocks were prepared after fixation of the tissues in periodate-lysine-paraformaldehyde buffer for 24 h and transfer to PBS. Sections 3 µm thick were prepared. Deparaffinized sections were processed for antigen retrieval by immersion in citric acid (pH 6.0) for 10 min and boiling in a microwave oven for 3 min. Immunohistochemical staining was performed using LSAB-2 kit (Dako, Carpenteria, CA).

Quantification of immunofluorescence was performed separately for cortex, outer medulla, and inner medulla because these areas have been shown to have varying expression of NOS and variable effects of L-NAME inhibition (45, 60). Scoring was performed by a previously described method (36). Intensity of staining for all antibodies was scored from 0 to 3, with "0" indicating absence of staining and "3" indicating maximal staining intensity. All preparations were evaluated by two independent investigators without knowledge of the source of renal sections.

In a separate series of experiments, immunofluorescence detection of endothelial and mesenchymal markers in the kidneys was performed with a laser confocal fluorescence microscopy and a Zeiss LSM 510 confocal microscope (Carl Zeiss, Thornwood, NY).

Assessment of vascular density. Scoring was performed per 100 glomeruli or respective tubular cross sections using a previously described method (8, 19). At least 100 fields sized 0.54 mm2 were examined in each area in at least four different animals per group. Additionally, the number of positively stained capillary cross sections was counted in at least 100 glomeruli for each group, and glomerular capillary density was expressed as a number of GFP-positive capillary sections per glomerulus.

Preparation of single cell suspensions from whole kidneys and fluorescence-activated cell sorter analysis. For preparation of single cell suspensions from whole kidneys, excised organs were placed on ice into culture dishes and cut into 1-mm3 pieces. Four milliliters of RPMI 1640 (Invitrogen) containing 1 mg/ml collagenase type II (Invitrogen) were added. The tissues were further minced in the solution, followed by incubation at 37°C in 5% CO2 for 30 min. Suspensions were filtered through a 38-µm steel mesh (W.S. Tyler). Repeated digestions for 10 min were optionally performed until microscopic investigation showed a single cell suspension. Further digestion was finally blocked by adding PBS with 1% (wt/vol) BSA, and cells were repeatedly washed for three times in PBS-BSA.

Quantification of the subpopulation of GFP-positive cells within the population of renal cells in suspension was performed by fluorescence-activated cell sorter (FACS) analysis. To define the correct gating parameters, kidneys from respective wild-type animals (FVB/NJ mice; Jackson Laboratories) were analyzed initially. Approximately 106 cells from every kidney were incubated for 30 min on ice with anti-{alpha}-SMA or anti-calponin antibodies in 1% PBS-BSA. Secondary antibodies used were Alexa fluor 594 donkey anti-mouse (Molecular Probes, Eugene, OR), with incubation performed for 30 min on ice. Thus processed cells were washed with 1% PBS-BSA and fixed in 4% paraformaldehyde. Data were acquired by a FACScan cytometer equipped with a 488-nm argon laser and a 635-nm red diode laser and analyzed using CellQuest software (Becton Dickinson, San Jose, CA). The setup of FACScan was achieved according to the manufacturer’s instructions using unstained cells.

Statistical analyses. Data are expressed as means ± SD. The means of two populations were compared by a Student’s t-test. To compare means of several populations, one-way ANOVA was used (unless indicated otherwise in RESULTS), followed by Tukey’s posttest for multiple comparisons. Differences were considered significant when P < 0.05.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
In vitro validation of L-NAME effect on collagen XVIII gene and protein expression in HUVEC. We considered one of the findings obtained through screening of cDNA microarray "cardiovascular-relevant" genes, namely, the upregulation of collagen XVIII, the precursor for endostatin, to be of potential importance. In these preliminary screenings for differential gene expression, the following set of data was obtained: gene = collagen XVIII; control = 32; L-NAME = 60; ratio = 1.9. Semiquantitative RT-PCR confirmed this finding: there was a significant increase in the ratio of collagen XVIII to 18S mRNA expression by HUVEC treated with L-NAME or ADMA (Fig. 1A; representative of 3 similar experiments; P < 0.05, multivariate ANOVA) compared with a vehicle. Western blot analysis of lysates obtained from cultured HUVEC demonstrated increased expression of endostatin fragment (22 kDa) of collagen XVIII after 24–48 h of treatment with either L-NAME or ADMA (Fig. 1B). To determine whether this phenomenon can be reproduced in animals receiving NOS inhibitor, additional in vivo studies were performed to validate the above findings.


Figure 1
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  		Fig. 1. Collagen XVIII expression in human umbilical vein endothelial cells (HUVEC). A: quantitative RT-PCR of collagen XVIII transcripts in HUVEC treated with PBS (vehicle control), asymmetric dimethylarginine (ADMA; 5 µM), and NG-nitro-L-arginine methyl ester (L-NAME; 50 µM). Results are representative of 3 different experiments. B: Western blot analysis of collagen XVIII abundance in HUVEC treated with PBS (vehicle control), ADMA (5 µM), and L-NAME (50 µM). Results are representative of 3 different experiments. Equal amounts of protein were used for loading of gels, and beta-actin expression was used to document the equal protein loading.

 
In vivo model of nonhypertensive chronic NOS inhibition. We sought to establish a nonhypertensive mouse model of NOS inhibition. The dose of 0.4 mg/ml L-NAME in drinking water was selected as the maximal nonhypertensinogenic dose. The level of blood pressure remained comparable in control and L-NAME groups throughout the study, except for the last measurement on week 7, when there was a small but significant increase in the systolic blood pressure in the 0.4 mg/ml L-NAME-treated group compared with controls (mean ± SD: 98 ± 21 in L-NAME vs. 85 ± 11 mmHg in control; P = 0.015); however, these values were well within the normal blood pressure range. There was no significant difference in the weight of animals or their kidneys. Protein-to-creatinine ratio (not shown) was increased in the L-NAME group compared with the vehicle or D-NAME-treated animals, but the difference failed to reach significance (P = 0.08).

To examine whether chronic 2-mo-long inhibition of NOS in vivo is accompanied by a similar upregulation of collagen XVIII/endostatin expression, immunohistochemical studies were performed and the intensity of staining was blindly scored. As illustrated in Fig. 2A, collagen XVIII/endostatin was barely detectable in control kidneys, whereas it was abundantly present in L-NAME-treated animals. Medullary accumulation of collagen XVIII/endostatin was remarkably and uniformly enhanced, whereas cortical accumulation showed a striped pattern along vascular bundles. Overall, summed scores indicated increased expression of collagen XVIII/endostatin (Fig. 2A), thus supporting the in vitro findings. Western blot analysis, which allowed us to separately analyze endostatin expression by its electrophoretic mobility, showed increased abundance of endostatin in the kidneys of mice chronically treated with L-NAME (Fig. 2B). Hence, increased expression of endostatin in endothelial cells and kidneys of mice subjected to pathophysiologically relevant concentration of ADMA (or to L-NAME in vivo) were confirmed at the level of mRNA and protein expression in vitro and protein expression and localization in vivo.


Figure 2
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  		Fig. 2. Expression of collagen XVIII and endostatin in Tie-2/green fluorescent protein (GFP) mice chronically treated with L-NAME or NG-nitro-D-arginine methyl ester (D-NAME). A: immunohistochemical staining of collagen XVIII/endostatin. Top: control kidney. Middle and bottom: kidneys from L-NAME-treated mice (cortex, glomerulus, papilla). B, top: scoring of the abundance of collagen XVIII in the cortex and medulla (in relative units). Note accumulation of this precursor of endostatin in both the cortex and medulla of L-NAME-treated mice. Data are means ± SD. B, middle: expression of endostatin in whole kidney lysates. Representative blots were obtained from individual animals; approximate molecular weights (MW, as measured in kDa) of the bands are shown at right. beta-Actin expression confirmed uniform gel loading. B, bottom: summary of the ratio of endostatin to beta-actin in control and L-NAME-treated mice. Data are means ± SD. *P < 0.05.

 
Profibrotic and antiangiogenic effects of L-NAME in vivo. Antiangiogenic actions of endostatin in endothelial cells have recently been explored on genomic and proteomic levels (1), but its potential profibrotic actions remain unexplored. Yet, there is indication that NO inhibition may upregulate expression of TGF-beta (34). Therefore, we used pan-TGF-beta antibody for immunohistochemical analyses and Western blotting of the kidneys. Immunohistochemical analysis showed that there was increased staining for a profibrotic marker, TGF-beta, in all zones of the renal parenchyma. Western blotting confirmed the increased expression of TGF-beta in L-NAME-treated animals (Fig. 3).


Figure 3
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  		Fig. 3. Expression of transforming growth factor (TGF)-beta in the kidneys of Tie-2/GFP mice chronically treated with L-NAME. Top: representative results of Western blot analysis of whole kidney lysates. C, control; L-N, L-NAME. Bottom: summary of ratio of TGF-beta to beta-actin expression in the kidney lysates. Data are means ± SD.

 
In view of the antiangiogenic activity of endostatin, we sought to investigate the possibility of microvascular rarefaction in the kidneys of mice with chronic inhibition of NOS. Capillary density of kidney sections was quantified by fluorescence microscopy of GFP as a marker of endothelial cells. (Examples of images used for such quantification are presented in GoFig. 5.) Figure 4 summarizes the data on capillary density in the cortex and medulla of kidneys of experimental animals. Statistically significant rarefaction of microvasculature was observed only in the inner medulla after 7 wk of treatment with nonhypertensinogenic doses of L-NAME. These data were in agreement with the scenario in which inhibition of eNOS resulted in the rarefaction of capillary profiles potentially leading to chronic ischemia, which in turn induced profibrotic changes.


Figure 4
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  		Fig. 4. Distribution of capillary density in the kidneys of Tie-2/GFP mice treated with L-NAME or D-NAME. GFP fluorescence was used as a guide for quantification of the number of endothelial cells per field (or per 100 glomerular or tubular profiles), as illustrated in Fig. 5. Data are means ± SD. Only the inner medulla showed a statistically significant difference between control and L-NAME groups (*P < 0.05).

 

Figure 5
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  		Fig. 5. Coexpression of {alpha}-smooth muscle actin ({alpha}-SMA) and GFP in the kidneys. Kidney cryosections were obtained from Tie-2/GFP mice and stained with antibodies to {alpha}-SMA. Images were obtained with green and red filters and merged. A: arteriolar blood vessel shows distinctive staining of the endothelium (green) and smooth muscle layer (red). Note complete separation of fluorescence of 2 fluorophores, indicative of the lack of "bleeding" of fluorescence emitted by each fluorophore to the images obtained with the second fluorophore. B: peritubular capillaries in a control kidney show GFP fluorescence devoid of {alpha}-SMA staining. C: glomeruli show GFP fluorescence that does not colocalize with {alpha}-SMA; D: in L-NAME-treated Tie-2/GFP mice, preponderance of peritubular capillaries displayed GFP and {alpha}-SMA fluorescence in a cell type-segregated fashion. E: a small proportion of capillaries showed GFP and {alpha}-SMA fluorescence confined to the same endothelial cell (orange-yellow on the merged images, arrows in F), as exemplified in a boxed area. Boxed area is shown enlarged in F.

 
In vivo detection of coexpression of markers of endothelial and mesenchymal cells. Capillary rarefaction results from either withdrawal of proangiogenic growth factors or accumulation of antiangiogenic substances (the latter scenario is applicable to our model). Either process could be expected to lead to the loss of endothelial differentiation markers, and these were studied in the next series of experiments. Fluorescence microscopy of GFP expression by endothelial cells in kidney cryosections was combined with immunofluorescence detection of {alpha}-SMA. As illustrated in Fig. 5, peritubular capillaries and glomeruli in control kidneys showed GFP fluorescence confined to the endothelium and unaccompanied by the {alpha}-SMA colocalization. In L-NAME-treated mice, the majority of peritubular capillaries also displayed GFP, whereas {alpha}-SMA fluorescence was confined to smooth muscle cells and myofibroblasts. However, a small proportion of capillaries showed GFP and {alpha}-SMA fluorescence confined to the same endothelial cell. The latter finding suggested that some endothelial cells were undergoing the process of transition from the endothelial to the mesenchymal phenotype.

Confocal microscopy confirmed these findings and revealed scattered individual cells within the peritubular capillaries, which coexpressed Tie-2/GFP and {alpha}-SMA. In addition, to mitigate any potential leakage of GFP from dysfunctional endothelial cells and its incorporation into nonendothelial neighboring cells, we examined the coexpression of mesenchymal ({alpha}-SMA) and endothelial (Tie-2) markers in the kidneys of mice deficient in eNOS and neuronal NOS using confocal fluorescence microscopy. The data showed a much more pronounced colocalization of these markers in the peritubular capillaries (Fig. 6). The fact that these models of genetic NOS ablation were characterized by a readily detectable colocalization of endothelial and mesenchymal markers argues in favor of NO deficiency as a leading mechanism for endothelial-mesenchymal transformation.


Figure 6
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  		Fig. 6. Laser scanning confocal microscopy imaging of the kidneys. Kidneys were obtained from mice lacking endothelial nitric oxide synthase (eNOS) and neuronal NOS (nNOS), stained with endothelial and mesenchymal markers, and compared with kidneys from Tie-2/GFP mice treated with L-NAME. Note that only merged images are presented. A: kidney section of L-NAME-treated Tie-2/GFP animal, staining for {alpha}-SMA. B: representative image of a kidney obtained from nNOS knockout mouse, double-staining for Tie-2 and {alpha}-SMA. C: representative image of a kidney obtained from eNOS knockout mouse, double stained for Tie-2 and {alpha}-SMA. Note that in merged images individual cells show colocalization of Tie-2 and {alpha}-SMA and that colocalization is much more prominent in nNOS and eNOS knockout animals than in Tie-2/GFP mice treated with L-NAME.

 
To verify the immunohistochemical findings using an independent approach, we next performed FACS analysis of collagenase-dispersed populations of kidney resident cells obtained from Tie-2/GFP mice, as detailed in MATERIALS AND METHODS. GFP-positive cells represented ~2% of the population. Staining of cells with antibodies against mesenchymal markers, calponin and {alpha}-SMA, showed a distinct subpopulation of cells, which exhibited an overlap with the GFP-expressing cells (Fig. 7A). The proportion of cells expressing both the endothelial and the mesenchymal markers was higher in kidneys from mice that received L-NAME (Fig. 7, B and C) than in controls. These findings further support the occurrence of endothelial-mesenchymal transformation induced by NO deficiency.


Figure 7
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  		Fig. 7. Fluorescence-activated cell sorter analysis of collagenase-dispersed kidney cells: coexpression of endothelial and mesenchymal markers. Cells were isolated and labeled as detailed in MATERIALS AND METHODS. A and B: representative examples of cell populations coexpressing Tie-2 promoter-driven GFP and {alpha}-SMA in control (A) and L-NAME-treated (B) mice. C: summary of fluorescence-activated cell sorter analyses (n = 3 animals each). Tie-2/GFP-labeled cells and {alpha}-SMA-positive cells represent each under 2% of total cell population obtained from collagenase-dispersed kidneys. The subpopulation of Tie-2/GFP-expressing cells costained with {alpha}-SMA is increased in L-NAME-treated mice (right). *P < 0.05. PE, phycoerythrine.

 
ADMA and endostatin induce the switch of cultured endothelial cells to the mesenchymal phenotype. The above demonstration of vascular rarefaction and occasional coexpression of {alpha}-SMA with GFP, driven by the endothelium-specific Tie-2 promoter, as well as colocalization of endothelial and mesencymal markers in microvasculature of eNOS knockout mice, suggested the possible participation of endothelial-mesenchymal transformation in the vasculopenia induced by the inhibition of eNOS. This possibility was addressed next with the use of in vitro studies seeking to simultaneously detect endothelial and mesenchymal markers in HUVEC. As shown in Figs. 8 and 9, ADMA treatment (5–15 µM for 48–72 h) resulted in a decreased expression of endothelial markers eNOS and Flk-1 with the concomitant increase in the expression of calponin, fibronectin, TGF-beta, and CTGF. ADMA effect could be reproduced by endostatin (100 nM for 48–72 h) or by TGF-beta2 (1 nM for 48–72 h) (Fig. 8). There was only a minimal change in the expression of CD31 or caveolin-1 (data not shown).


Figure 8
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  		Fig. 8. ADMA-induced dose-dependent loss of Tie-2 and gain of {alpha}-SMA by endothelial cells. A: representative images of HUVEC treated with 5 and 15 µM ADMA with immunocytochemical detection of endothelial marker Tie-2. Magnification = x630. Right: summary of fluorescence intensity. au, Arbitrary unit. B: representative images of HUVEC treated with 5 and 15 µM ADMA with immunocytochemical detection of mesenchymal marker {alpha}-SMA. Magnification = x630. Right: summary of fluorescence intensities. asm, {alpha}-SMA. Data are means ± SD. *P < 0.05.

 

Figure 9
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  		Fig. 9. Coexpression of endothelial and mesenchymal markers in endothelial cells exposed to ADMA, endostatin, or TGF-beta. Experiments were performed as in Fig. 8, except that additional groups of cells were treated with endostatin or TGF-beta. A: representative results of Western blotting. CTGF, connective tissue growth factor. B: summary diagrams for ratios of eNOS to beta-actin and calponin to beta-actin expression. Data are means ± SD. *P < 0.05 vs. control (Contr).

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
This study presents two sets of major findings. The first ascribes to NO deficiency in endothelial cells the function of a profibrotic stimulus associated with the expression of an antiangiogenic fragment of collagen XVIII, endostatin. The second set provides evidence of endothelial-mesenchymal transdifferentiation in the course of inhibition of NOS by a pathophysiologically important antagonist, ADMA. Both could account for the microvascular rarefaction as a forerunner of tubulointerstitial fibrosis. The concept of endothelial cell dysfunction triggering profibrotic changes in the kidney, as it stems from the results of our studies, is not entirely novel. Nagata et al. (43) observed ultrastructural changes in glomerular endothelial cells at sites destined to develop glomerulosclerosis after unilateral nephrectomy. In rats with subtotal 5/6 nephrectomy, glomerular endothelial dysfunction was associated with increased local expression of mRNA for angiotensinogen, TGF-beta1, fibronectin, and laminin before development of glomerulosclerosis (34). A possible role of impaired NO synthesis in the fibrogenic transformation has been suggested. Hogaboam et al. (22) convincingly demonstrated that pulmonary granulomas (induced by purified protein derivative, a model of nonfibrotic inflammatory process) in mice receiving L-NAME or aminoguanidine inhibitors of NOS were larger in diameter and had higher collagen I content than animals with intact NOS function. In cultured mesangial cells, elevated ambient glucose level resulted in the induction of TGF-beta mRNA and bioactivity accompanied by enhanced collagen synthesis (10), whereas exogenous or endogenous NO suppressed the induction of TGF-beta and collagen synthesis. Cocultured endothelial and vascular smooth muscle cells exhibited NO-dependent deposition of collagen I (42). Likewise, inhibition of NOS with L-NAME resulted in increased extracellular matrix deposition and induction of collagen I mRNA and accumulation of collagen I in the afferent arterioles and glomeruli of mice (9). Collectively, these data suggest the existence of a link between NO production and fibrotic transformation in different tissues.

The data presented herein further implicate NO deficiency in the development of a profibrotic phenotype of endothelial cells. The following lines of experimental evidence support this conclusion: 1) results of cDNA microarray survey of genes modulated by NO deficiency in HUVEC, namely, upregulation of collagen XVIII; 2) confirmation of this finding in vitro and in vivo; 3) occurrence of profibrotic changes in mice treated with nonhypertensinogenic doses of L-NAME, that is, increased expression of TGF-beta and CTGF; and 4) its association with the rarefaction of capillary profiles. Importantly, the observed in vivo renal profibrotic changes occurred in the absence of hypertension or any interventions required for blood pressure control. This distinct characteristic of our study offers a highly plausible insight into the independent role of NO deficiency and developing endothelial dysfunction in the regression of capillaries and profibrotic changes.

Significant literature exists on the role of epithelial-mesenchymal transformation in development and progression of fibrosis (24, 26, 37, 54). However, the role of endothelial-mesenchymal transformation in vascular regression and fibrosis remains unexplored. In a recent study of an accelerated model of ANG II-induced renal fibrosis, expanding interstitial cells expressing {alpha}-SMA originated from the perivascular spaces (15), rather than from epithelial-mesenchymal transformation. The data presented here are consistent with the contributory role of endothelial-mesenchymal transformation. It has been appreciated that, during embryonic development, Flk-1-positive vascular progenitor cells may differentiate into either endothelial or smooth muscle cells (56). DeRuiter et al. (11) documented the potential of embryonic endothelial cells to transdifferentiate into mesenchymal cells expressing smooth muscle cell markers. These observations posed a broader question on the potential single progenitor cell type for endothelial and smooth muscle cells and the possibility of conversion of endothelial to smooth muscle cells during adulthood. In vitro studies of cultured bovine and human endothelial cells showed that gamma-irradiation, mitomycin C, or FGF deprivation induces endothelial transformation into smooth muscle cells (18, 23). Our data demonstrate that ADMA induces the loss of endothelial differentiation markers, such as eNOS or Flk-1, while increasing the expression of mesenchymal markers, such as calponin or fibronectin. Inhibition of NO synthesis also led to the increased expression of collagen XVIII/endostatin, TGF-beta, and CTGF in cultured endothelial cells and in the nonhypertensive animal model. Exposure of cultured endothelial cells to either endostatin or TGF-beta mimicked the effect of ADMA, suggesting their role in effecting endothelial-mesenchymal transformation. In turn, enhanced expression of TGF-beta in HUVEC was achieved by both ADMA and endostatin. This may be due to the existence of two independent pathways (NO deficiency mediated and endostatin mediated), both triggered by ADMA, in switching the endothelial phenotype toward the mesenchymal. Alternatively, this could be explained by the ability of endostatin to upregulate TGF-beta. Additional profibrotic pathogenetic mechanisms of collagen XVIII/endostatin accumulation may be governed by its recently recognized role in the epithelial-mesenchymal transformation in the atrioventricular valves during cardiac development (7) and/or interaction with L-selectin and monocyte chemoattractant protein-1 (31), thus contributing to leukocyte infiltration and renal inflammation.

The described role of collagen XVIII/endostatin in profibrotic changes may well be expanded beyond the kidney. In a study of fibrotic and cirrhotic human livers, collagen XVIII was found to be overexpressed in hepatocytes and in endothelial and bile duct cells (41). In patients with ischemic cerebral events, elevated endostatin levels were independently associated with the higher risk of recurrence and progression of intracranial atherosclerosis (2). Furthermore, in mice kept under conditions of hypobaric hypoxia, elevated expression of endostatin occurred in lungs and aorta, contributing to development of pulmonary hypertension as a result of impaired angiogenesis (49). These multidisciplinary findings attest to the potentially broader contribution of collagen XVIII/endostatin in the development of fibrosis.

It is plausible that the capillary rarefaction, most prominent in the renal medulla, is a consequence of overproduction of endostatin with its antiangiogenic properties (48). Studies of a recently engineered collagen XVIII-deficient mouse reconfirmed the role of this COOH-terminal fragment in antiangiogenesis: angiogenesis was significantly enhanced in explant aortic cultures obtained from knockout mice compared with wild-type animals (36). Our data may offer a link between the two categories of profibrotic action of endostatin: 1) a direct one due to the phenotypical switch of endothelial cells and 2) an indirect one due to the vascular rarefaction and developing renal hypoxia. Specifically, NO inhibition and overexpression of endostatin and TGF-beta could actuate endothelial-mesenchymal transdifferentiation, which in turn participates in vascular rarefaction. Clearly, the mechanisms of NO regulation of fibrogenesis are far from being resolved. Nonetheless, the data presented here provide support to the idea that endothelial dysfunction, as exemplified by the inhibition of NOS, provokes profibrotic phenotypical switch in endothelial cells and triggers their mesenchymal transformation (20, 27–30, 35, 46, 47).


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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REFERENCES
 
These studies were supported in part by National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-45462, DK-54602, DK-52783 (M. S. Goligorsky), National Kidney Foundation award (E. O’Riordan), and Westchester Artificial Kidney Foundation.


   FOOTNOTES
 

Address for reprint requests and other correspondence: M. S. Goligorsky, New York Medical College, Basic Sciences Bldg., Rm. C23, Valhalla, NY 10595 (e-mail: michael_goligorsky{at}nymc.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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