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From the * Andrology Unit, Department of
Physiopathology, Center for Research, Transfer and High Education DENOTHE, the
Department of Anatomy, Histology and Forensic
Medicine, and the Interdepartmental Laboratory
of Functional and Cellular Pharmacology of Reproduction, Departments of
Pharmacology and Clinical Physiopathology, University of Florence, Florence,
Italy; and Bioxell, Milan, Italy.
| Correspondence to: Dr Mario Maggi, Andrology Unit, Department of Clinical Physiopathology, University of Florence, V.le G. Pieraccini, 6, 50139 Florence, Italy (e-mail: m.maggi{at}dfc.unifi.it). |
| Received for publication April 23, 2007; accepted for publication August 9, 2007. |
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Abstract |
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-smooth muscle actin, SM22, and myocardin. In contrast to
atorvastatin, elocalcitol, a vitamin D analog that also interferes with RhoA
activation in SHR bladder, was unable to restore penile responsiveness to
sildenafil. In conclusion, atorvastatin, but not elocalcitol, ameliorates
sildenafil-induced penile erections in SHR, likely by interfering with
RhoA/ROCK signaling within the penis.
Key words: Erectile dysfunction, phosphodiesterase 5 inhibitor, human penile smooth muscle cells, RhoA-dependent genes, SHR model
RhoA is a member of a small monomeric GTPase family that is involved in SM
contraction and the regulation of several other SM-dependent processes such as
cell adhesion (Ren et al,
1999), motility (Wheeler and
Ridley, 2004), migration
(Wheeler and Ridley, 2004),
and proliferation (Aznar and Lacal,
2001). As soon as ET1 and NA bind to their excitatory receptors,
RhoA is converted from the cytoplasmic, inactive GDP-bound form into an active
GTP-bound complex that translocates to the plasma membrane, where it binds via
geranylgeranylation, initiating signal transduction. The best-characterized
downstream effectors of RhoA are Rho-associated, coiled-coil–containing
protein kinases (ROCKs), which are directly involved in SM contraction
(Noma et al, 2006). The 2
described ROCK isoforms (ROCK1 or ROKβ and ROCK2 or ROK), sharing
65% homology in amino acid sequence and 92% homology in their kinase domains,
are serine-threonine kinases that are able to maintain the phosphorylated
state of the myosin light chain and thus the contractile tone independently of
intracellular calcium levels. More recently the RhoA/ROCK pathway has been
involved in the "excitation-transcription coupling" of SM cells
(Barlow et al, 2006). RhoA/ROCK
signaling stimulates the transcription of SM-specific genes, such as
-SM actin (-SMA), SM myosin heavy chain, and desmin, through the
induction of myocardin, a transcriptional coactivator controlling several
genes involved in SM commitment and function
(Wamhoff et al, 2004;
Pipes et al, 2006).
RhoA/ROCK pathway overactivity has been shown to be involved in several pathophysiologic processes such as angiogenesis, atherosclerosis, cerebral and coronary vasospasm, cerebral ischemia, glomerulosclerosis, hypertension, myocardial hypertrophy, myocardial ischemia-reperfusion injury, neointima formation, pulmonary hypertension, vascular remodeling, diabetes mellitus, and ED (Chrissobolis and Sobey, 2006; Jin and Burnett, 2006; Moore and Wang, 2006; Noma et al, 2006; Calò and Pessina, 2007). ED is often associated with hypertension (Billups et al, 2005; Mulhall et al, 2006), and both conditions might share similar pathogenetic determinants. A longitudinal study in spontaneously hypertensive rats (SHR) clearly showed that changes in erectile tissue precede aortic ones (Behr-Roussel et al, 2006), as suggested also in humans (Billups et al, 2005; Montorsi et al, 2006). RhoA is overactive and hypersensitive to G protein–coupled receptor stimulation in several SHR tissues, including vascular SM cells (Moriki et al, 2004; Ying et al, 2004), brainstem (Sagara et al, 2007), and preglomerular microvascular SM cells (Jackson et al, 2005). It has been postulated that RhoA overactivity significantly contributes to the development of hypertension in this rat strain. A similar finding was also reported in the penis (Wilkes et al, 2004), where the up-regulation of RhoA/ROCK signaling is considered to play an important role in hypertension-associated ED (Chitaley et al, 2001; Wilkes et al, 2004). Accordingly, Y-27632 administration partially restores erectile function (Wilkes et al, 2004) and synergizes with phosphodiesterase 5 (PDE5) inhibition through zaprinast (Chitaley et al, 2001).
The recent introduction of the PDE5 inhibitor sildenafil has greatly improved the success rate of ED treatment; however, hypertensive patients are usually less responsive to sildenafil than normotensive ones (Chia et al, 2004). The aim of this study was to systematically investigate the in vivo effects in SHR compared with their normotensive counterpart, Wistar-Kyoto (WKY) rats, sildenafil treatment as monotherapy or in combination with atorvastatin, a drug known to interfere with the RhoA/ROCK pathway by decreasing RhoA geranylgeranylation (Bonetti et al, 2003; Rikitake and Liao, 2005). The effects of atorvastatin were compared with those obtained with elocalcitol (also known as BXL-628), a nonhypercalcemic vitamin D analog that has been recently demonstrated to interfere with RhoA activation in SHR bladder, characterized by RhoA/ROCK overactivity (Morelli et al, 2007). The mechanism by which atorvastatin is able to interfere with RhoA activity has been studied using a previously characterized cellular model of human penile SM cells (hfPSMC; Granchi et al, 2002; Crescioli et al, 2003b; Filippi et al, 2003a,b; Vignozzi et al, 2005).
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Methods |
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Rat Tissues
Rats were sacrificed by cervical dislocation. Urinary bladders and CC were
removed and immediately frozen in liquid nitrogen at –80°C until RNA
preparation. Experiments were performed in accordance to the Italian
Ministerial Law 116/92 and approved by the Institutional Animal Care and Use
Committee of the University of Florence, Florence, Italy.
Cholesterol and Calcium Measurements
Blood samples for total cholesterol measurements were obtained at baseline
via the femoral veins. The blood was immediately centrifuged at
1000xg for 20 minutes, and the plasma was stored at
–20°C until analyzed. Plasma cholesterol levels were measured with
an automated system (Abbott Diagnostics, Abbott Park, Ill). Serum calcium
levels were measured with a commercially available colorimetric assay
(Sigma-Aldrich, St Louis, Mo) according to the manufacturer's instructions and
as previously reported (Crescioli et al,
2003a).
Human Tissues
Human tissues (testis, epididymis, CC, prostate, bladder, heart, brain, and
skeletal muscle) were collected during surgery for benign diseases. All tissue
samples were obtained after the approval of the Hospital Committee for
Investigation in Humans (protocol 6783-04; Azienda Ospedaliera Universitaria
Careggi, Florence, Italy) and after receiving consent from the informed
patients. Immediately after removal, tissue samples were shock-frozen in
liquid nitrogen and stored at –80°C until RNA preparation.
CC Cell Cultures
hfPSMC were prepared from 7 samples of fetal male external genitalia
(11–12 weeks of gestation) obtained after spontaneous or therapeutic
abortion, as previously described (Granchi
et al, 2002; Crescioli et al,
2003a; Filippi et al,
2003a,b;
Vignozzi et al, 2005). The
experiments were performed using hfPSMC from the seventh to the 13th passage
in Dulbecco modified Eagle medium supplemented with 5 mM glucose and Ham F12
medium.
Confocal Laser Microscopy
hfPSMC were seeded in growth medium onto sterile glass slides
(approximately 104 cells/mL) and incubated for 48 hours with ET1
(100 nM; Calbiochem, San Diego, Calif), atorvastatin (1 µM), or ET1 (100
nM) in combination with atorvastatin (1 µM). Cells in serum-free growth
medium were used as controls. Cells were then fixed with 3.7% paraformaldehyde
(pH 7.4) for 10 minutes, permeabilized for 10 minutes with phosphate-buffered
saline (PBS) containing 0.1% Triton X-100, and incubated with 2% bovine serum
albumin (BSA) for 15 minutes. Immunostaining was performed as previously
described (Morelli et al,
2007) using anti–pan-cadherin (1:500; Abcam Ltd, Cambridge,
United Kingdom) and anti-RhoA (1:100; Santa Cruz Biotechnology, Santa Cruz,
Calif) antibodies, followed by rhodamine red goat anti-mouse immunoglobulin
(IgG) heavy and light chains (H+L) (R6393; 1:200; Molecular Probes, Eugene,
Ore) and Alexa Fluor 488 goat anti-mouse IgG (H+L) (A11001; 1:200; Molecular
Probes) antibodies, respectively.
Preparation of Membrane/Cytosolic Fractions and Sodium Dodecyl Sulphate Polyacrylamide Gel Electrophoresis/Western Blot Analysis
Subconfluent hfPSMC were incubated for 24 hours in serum-free medium before
exposure to the following 48-hour treatments: ET1 (100 nM) in the absence or
presence of atorvastatin (1 µM) and atorvastatin alone (1 µM) or in
combination with geranylgeranyl pyrophosphate (GGPP; 1 µM; Sigma-Aldrich)
added 6 hours before cells were collected. Cells were collected using
trypsin-EDTA and were divided into 2 aliquots: one for total protein
extraction and the other for membrane/cytosolic preparations. Membrane and
cytosolic fractions were prepared using the ProteoExtract subcellular proteome
extraction kit (Calbiochem) according to the manufacturer's instructions.
Protein extracts were quantified with the BCA reagent (Pierce, Rockford, Ill).
Equal volumes of samples (15 µg) were resolved by 12% sodium dodecyl
sulphate polyacrylamide gel electrophoresis. Western blot analysis with an
anti-RhoA antibody (1:500; Santa Cruz Biotechnology) was performed as
previously described (Morelli et al,
2007). Equal protein loading was verified by Ponceau-S staining
(Sigma-Aldrich). Densitometric analysis of band intensity acquired by a plot
bed scanner (IS440CF; Kodak Digital Science, Cinisello Balsamo, Italy) was
performed using Photoshop 5.5 software (Adobe Systems Inc Italia srl, Agrate
Brianza, Italy).
ROCK Activity Assay
After serum deprivation for 24 hours, hfPSMC were stimulated for 48 hours
with ET1 (100 nM; Calbiochem) in the absence or presence of atorvastatin (1
µM). Cells in serum-free growth medium were used as controls. Cells were
harvested by rubber policeman using lysis buffer (50 mM Tris-HCl [pH 7.5], 1
mM EDTA, 1 mM EGTA, 0.4 mM paraphenyl methane sulphonyl fluoride, 1 µg/mL
pepstatin, 0.5 µg/mL leupeptin, 2 mM NaF, 2 mM
Na3VO4). Cells were lysed by 3 cycles of sonication.
Protein extracts were quantified with the BCA reagent (Pierce). Samples were
prepared for the kinase assay by immunoprecipitating equal amounts of cell
lysate with an anti-ROCK antibody (SC17794; Santa Cruz Biotechnology), as
previously described (Morelli et al,
2007). The immunokinase assay was carried out using the CycLex
Rho-kinase Assay kit (MBL International, Way Woburn, Mass) following the
manufacturer's instructions.
Cell Proliferation Assay
hfPSMC (2.3 x 104) were seeded into 12-well plates in growth
medium. After 24 hours, the cells were washed in PBS and incubated overnight
in phenol red and serum-free medium containing 0.1% BSA. In the first set of
experiments, increasing concentrations of atorvastatin (10 nM–100 µM)
or elocalcitol (10 pM–100 nM) were added for 48 hours together with a
fixed concentration of platelet-derived growth factor BB (PDGF-BB; 25 ng/mL;
Sigma-Aldrich). In a second set of experiments, the effect of atorvastatin (1
µM) on PDGF-BB–induced cell growth was tested in the presence or
absence of the selective RhoA inhibitor C3 exoenzyme (1 µg/mL; Calbiochem),
guanylate cyclase inhibitor
1H-[1,2,4]oxadiazolo[4,3-]quinoxalin-1-one (ODQ; 1 µM;
Tocris, Bristol, United Kingdom), or nitric oxide inhibitor
NG-nitro-L-arginine methyl ester (L-NAME; 1 µM;
Sigma-Aldrich). In all experiments, PDGF-BB (25 ng/mL) was also tested alone,
and untreated cells in serum-free growth medium were used as controls. After
48 hours, cells were trypsinized, and the proliferation rate was derived by
hemocytometer counting of 9 different fields for each well. Each experimental
point was repeated at least in triplicate in at least 3 different experiments.
Results are expressed as the percent variation (mean ± SEM) over
PDGF-BB or basal conditions.
Cell Chemotaxis Assay
The effect of increasing concentrations of elocalcitol (10 pM–100 nM)
or atorvastatin (10 nM–100 µM) on ET1-induced hfPSMC migration was
studied in a P48 multiwell Boyden chamber (Nuclepore Inc, Pleasanton, Calif).
Chemotaxis was evaluated as previously described
(Vignozzi et al, 2005;
Morelli et al, 2007) using
polyvinyl-pyrrolidine–free polycarbonate filters with an 8-µm pore
size coated with 20 µg/mL type I collagen (BD Biosciences, Bedford, Mass).
ET1 (100 nM in 28 µL) was added to the lower wells, and cells (1 x
104 cells in 40 µL) were seeded into the upper wells of the
chamber with or without elocalcitol or atorvastatin and incubated at 37°C
for 5 hours. The effects of atorvastatin (1 µM) in the presence of
increasing concentrations (10 nM–1 µM) of GGPP (Sigma-Aldrich) or
elocalcitol (10 nM) in the presence of 1 µM GGPP were also tested on
ET1-induced (100 nM) cell migration. Unstimulated cells in serum-free culture
medium were used as controls of basal migration. The inhibitory effect on
ET1-induced migration was tested using methanol-fixed cells stained with
Diff-Quick (DADE Behring AG, Düdingen, Switzerland), and cell migration
was measured by microscopic evaluation of the number of cells that moved
across the filter into 10 random fields. Each experimental point was
replicated at least 5 times. Results were obtained from 3 independent
experiments. Data are expressed as mean ± SEM of the percentage
increase with ET1-induced migration or basal conditions as 100%.
Real-Time Quantitative Reverse Transcription Polymerase Chain Reaction
Isolation of RNA from rat and human tissues and hfPSMC cells was performed
using the RNAeasy kit (QIAGEN, Valencia, Calif). cDNA synthesis was performed
as previously described (Morelli et al,
2007). mRNA quantitative analysis was performed according to the
fluorescent TaqMan methodology (Morelli et
al, 2004; Morelli et al,
2007). Polymerase chain reaction (PCR) primers and probes specific
for mRNA sequences of target genes (RhoA, ROCK1, ROCK2, endothelial
nitric oxide synthase [eNOS], neuronal NOS [nNOS],
PDE5, vitamin D receptor [VDR], desmin, -SMA,
transgelin or SM22, and myocardin) were purchased from Applied
Biosystems (Foster City, Calif). β2-microglobulin and 18S rRNA
were chosen as reference genes for rat and human samples, respectively, and
were selected among the endogenous controls provided by Applied Biosystems.
The PCR mixture (25-µl final volume) consisted of 1X final concentrations
of primers and probe mix, 1X final concentration of a universal PCR master mix
(Applied Biosystems), and 25 ng of cDNA. Amplification and detection were
performed with the ABI Prism 7700 Sequence Detection System, as previously
reported (Morelli et al,
2007). Each measurement was carried out in duplicate. Data
analysis was based on the comparative Ct method according to the
manufacturer's instructions (Applied Biosystems).
Statistical Analysis
Results are expressed as mean ± SEM for n experiments. Statistical
analysis was performed with 1-way analysis of variance test followed by
Tukey-Kramer post hoc analysis, and P < .05 was considered
significant. Half-maximal response effective dose (ED50) and
half-maximal response inhibitory concentration (IC50) values were
calculated using the computer program ALLFIT
(De Lean et al, 1978).
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Results |
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Gene Expression Analysis in Penile Tissue From Experimental Rats
Expression of RhoA and its downstream kinases (ROCK1 and
ROCK2) in the different experimental groups, as evaluated by
real-time quantitative reverse transcription (RT) PCR (qRT-PCR), is shown in
Figure 2. mRNA expression of
RhoA, ROCK1, and ROCK2 was higher in CC samples from
untreated SHR compared with WKY rats. Elocalcitol did not revert this
overexpression. At both doses, atorvastatin treatment reduced RhoA
gene expression (Figure 2A),
whereas ROCK1 mRNA was unaffected
(Figure 2B). ROCK2
mRNA expression was inhibited by atorvastatin in a dose-dependent manner
(Figure 2C).
Atorvastatin-induced ROCK2 down-regulation resulted in statistically
significance (P < .05) only at the highest dose tested (30 mg/kg;
Figure 2C). Other genes
investigated, including nNOS, eNOS, and PDE5, were
unaffected by hypertensive conditions or atorvastatin treatment
(Table 1).
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Based on our previous observation that elocalcitol treatment down-regulates RhoA/ROCK overactivity in the bladder of SHR and reduces hypersensitivity to Y-27632 up to WKY levels (Morelli et al, 2007), we studied the expression of the elocalcitol receptor, VDR, in the bladders and CC of these rat strains. Interestingly, VDR expression was approximately threefold higher in the bladder, but not in the CC, of SHR compared with WKY rats (Table 2).
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Effect of Atorvastatin and Elocalcitol on the RhoA/ROCK Pathway in hfPSMC
To further investigate atorvastatin effects on the RhoA/ROCK pathway, we
used a previously characterized SM cell preparation from hfPSMC
(Granchi et al, 2002;
Crescioli et al, 2003b;
Filippi et al,
2003a,b;
Vignozzi et al, 2005).
Figure 3 shows the relative
mRNA expression of RhoA and its associated kinases, ROCK1
and ROCK2, in hfPSMC and the distribution of these transcripts in
several adult human tissues, including adult CC, as assessed by qRT-PCR.
RhoA mRNA is abundantly expressed in all the tissues studied, and of
the two RhoA-associated kinases, ROCK2 is the predominant isoform.
Because hfPSMC express high levels of RhoA and its associated kinases, these
cells represent a useful model to study the pharmacologic regulation of this
protein family.
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To study the effects of atorvastatin on RhoA membrane translocation, we used immunofluorescence confocal microscopy, as previously described (Morelli et al, 2007). We monitored the intracellular localization of RhoA in hfPSMC under basal conditions (Figure 4A) and after a 48-hour stimulation with ET1 (Figure 4D), atorvastatin (Figure 4G), or ET1 and atorvastatin (Figure 4L). Pan-cadherin was used as a cell membrane marker (Figure 4B, E, H, and M). ET1 increased RhoA localization at the plasma membrane, as evidenced by merging RhoA and pan-cadherin staining (Figure 4F). Atorvastatin alone (Figure 4I), and more impressively after ET1 stimulation (Figure 4N), reduced membrane translocation of RhoA, as shown in the merged images of Figure 4. To further validate these results, we studied by immunoblot the subcellular distribution of RhoA (membrane vs cytosol) and compared it with total RhoA. ET1 (100 nM) stimulated membrane expression of RhoA, which was reduced by atorvastatin (1 µM; Figure 5A, upper panel). Interestingly, atorvastatin treatment induced a robust RhoA accumulation in the cytosolic fraction (Figure 5A, middle panel). Figure 5B shows the effect of atorvastatin, with or without the addition of GGPP, on membrane-bound RhoA in multiple experiments (n = 3). As observed previously, atorvastatin (1 µM) significantly inhibited RhoA membrane translocation (P < .0001; Figure 5B), whereas the addition of GGPP (1 µM), by restoring RhoA geranylgeranylation, reverted the atorvastatin effects (P < .01) and increased the quantity of membrane-bound RhoA over the control level (P < .05; Figure 5B). Finally, to evaluate whether the activity of ROCK, the main kinase downstream of RhoA, was affected by atorvastatin treatment, we performed an immunokinase assay in hfPSMC (Figure 5C–E). In hfPSMC, ROCK activity was inhibited in time- and dose-dependent manners (Figure 5C) by the specific ROCK antagonist H-1152 (IC50 = 7.5 ± 0.4; n = 2; Figure 5D). Whereas ET1 (100 nM) stimulated a significant increase in ROCK activity (P < .05), atorvastatin (1 µM) completely prevented such an increase (n = 3).
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To investigate whether atorvastatin-induced reduction of RhoA membrane translocation affected RhoA-mediated cellular functions, we studied the effect of increasing atorvastatin concentrations on cell migration and proliferation of hfPSMC stimulated by ET1 (100 nM) or PDGF-BB (25 ng/mL), respectively. ET1 induced a 238% ± 17% (P < .0001) increase in cell migration, and PDGF-BB increased cell proliferation by 168% ± 7% (P < .0001). Atorvastatin treatment resulted in dose-dependent inhibition of both cell migration and proliferation, with a similar IC50 (shared IC50 = 0.97 ± 0.3 µM; Figure 6A). Inhibition of hfPSMC cell growth and motility was also induced by another inhibitor of RhoA membrane translocation, elocalcitol (Figure 6B). Again, IC50s for elocalcitol-induced inhibitory effects were similar (shared IC50 = 9.7 ± 3 nM). Addition of increasing concentrations of GGPP (10 nM–1 µM) reversed atorvastatin-induced inhibition of migration in a dose-dependent manner (Figure 6C). Conversely, the maximal concentration of GGPP (1 µM) was unable to revert elocalcitol activity (Figure 6C). Blocking RhoA activation with C3 exoenzyme, through ADP ribosylation, induced an anti-proliferative effect similar to that observed with atorvastatin (Figure 6D). Atorvastatin and C3 exoenzyme cotreatment did not enhance growth inhibition compared with either agent alone (Figure 6D). Blocking nitric oxide formation with the NOS inhibitor L-NAME (1 µM) or signaling with the guanylate cyclase inhibitor ODQ (1 µM) did not impair the antiproliferative action of atorvastatin (Figure 5D).
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-SMA, SM22, and myocardin
(Figure 7D–G).
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Discussion |
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Statins reduce cholesterol levels through a mevalonic acid–like
moiety that competitively inhibits 3-hydroxy-3-methylglutaryl coenzyme A
(HMG-CoA), the rate-limiting step in cholesterol biosynthesis. However,
independently of intracellular cholesterol biosynthesis, statins, through
competition for HMG-CoA, also inhibit the formation of isoprenoid
intermediates. These intermediates are essential for geranylgeranylation of
small GTP-binding proteins, including RhoA, and for their binding to the
plasma membrane, which is important for GDP-GTP exchange
(Bonetti et al, 2003;
Chrissobolis and Sobey, 2006).
In this study, we showed that in isolated hfPSMC
(Granchi et al, 2002;
Crescioli et al, 2003b;
Filippi et al,
2003a,b;
Vignozzi et al, 2005),
atorvastatin decreases ET1-induced RhoA membrane translocation and therefore
the downstream activation of ROCK and of the cascade leading to contractility,
motility, proliferation, and SM differentiation. Accordingly, in isolated
human penile cells, ET1-stimulated cell migration and PDGF-BB–induced
cell proliferation were inhibited by atorvastatin in a dose-dependent manner
at submicromolar concentrations. In addition, gene expression of the
predominant RhoA-activated kinase in erectile tissue, ROCK2, but not
ROCK1, was inhibited in SHR, as well as in hfPSMC. Specific SM genes
downstream of the RhoA/ROCK cascade, such as desmin, -SMA, SM22,
and myocardin, were also inhibited by atorvastatin treatment. It has been
previously demonstrated that overexpression of constitutively active RhoA
stimulates transcription of multiple CArG-dependent SM genes
(Mack et al, 2001;
Gudi et al, 2002), including
desmin and -SMA (Mericskay et al,
2000), or their coactivators, such as myocardin
(Wamhoff et al, 2004), and
that C3 exoenzyme, an inhibitor of RhoA activation, or Y-27632, an inhibitor
of ROCK activity, counteracts this induction. Hence, our data indicated that
atorvastatin can interact with the well-described
"excitation-contraction coupling" mediated by the RhoA/ROCK
pathway and interferes with the RhoA/ROCK-mediated
"excitation-transcription coupling" that regulates SM cell
commitment (Barlow et al, 2006;
Posern and Treisman,
2006).
Statins are known to increase NO bioavailability through several mechanisms, including stabilization of eNOS mRNA (Laufs et al, 1997, 1998), increases in eNOS phosphorylation by protein kinase B (Kureishi et al, 2000), or inhibition of caveolin 1 expression (Feron et al, 2001). Because we previously demonstrated that SM cells express a functional NOS (Filippi et al, 2003b) and that 2 distinct NO donors inhibit hfPSMC proliferation through cGMP formation (Filippi et al, 2003a), we hypothesized that increased NO/cGMP availability contributed to the atorvastatin-induced antiproliferative effect. Our results indicates that inhibition of NO formation (L-NAME) or action (ODQ) fail to affect the antiproliferative effect of atorvastatin. In addition, studies in rat CC fail to support a modulation of atorvastatin on gene expression of enzymes involved in cGMP generation (NOS) or degradation (PDE5). Incubation with C3 exoenzyme mimicked atorvastatin-induced growth arrest, further suggesting a direct effect of atorvastatin on RhoA activation, most probably by interfering with its geranylgeranylation. This view is supported by the observation that in the presence of GGPP, the effect of atorvastatin on RhoA membrane translocation and cell proliferation was completely reverted. In contrast, elocalcitol-induced growth arrest was not prevented by GGPP, suggesting that RhoA inhibition by this drug involves different mechanisms that remain to be investigated.
Our data confirm a previous observation of increased RhoA expression in penile tissue from SHR (Wilkes et al, 2004) and provide evidence that also the RhoA downstream kinases, ROCK1 and ROCK2, are up-regulated in SHR CC, as previously reported in systemic vasculature (Mukai et al, 2001). Interestingly, chronic atorvastatin dosing, at both concentrations tested, decreased RhoA mRNA expression up to WKY levels. Because atorvastatin does not directly change RhoA gene expression in isolated penile SM cells, we hypothesized that the RhoA penile changes observed in vivo in SHR might be due to more complex and indirect interactions involving cGMP formation or degradation. However, as mentioned before, we could not find any alteration in the cGMP-related genes investigated, including NOS isoform expression and PDE5. Because atorvastatin also increases NOS signaling through posttranscriptional mechanisms (Laufs et al, 1997; Laufs et al, 1998; Kureishi et al, 2000; Feron et al, 2001), it is still possible that RhoA down-regulation is cGMP mediated via NO release, as previously described (Chitaley et al, 2001; Lee et al, 2004). Accordingly, in the rat, an NO donor, NOR-1, and a ROCK inhibitor, Y-27632, synergize to induce in vivo penile erection (Mills et al, 2002), and endothelium denudation decreases Y-27632–induced relaxation in isolated rabbit CC (Filippi et al, 2003a,b).
In line with previous studies (Chitaley et al, 2001; Ushiyama et al, 2004; Wilkes et al, 2004; Behr-Roussel et al, 2006; Jin and Burnett, 2006), we found that erectile function in SHR was strongly compromised and responsiveness to sildenafil, although present, was relatively limited and lower than in their normotensive counterpart WKY rats. A key result of this study is that atorvastatin ameliorates sildenafil-induced erections in a dose-dependent manner in SHR. In particular, atorvastatin restored sildenafil responsiveness up to WKY levels, at least at maximal stimulation frequencies (16–32 Hz). Conversely, atorvastatin was unable to affect the ICP:MAP ratio in SHR not supplemented with sildenafil.
Our results are also in line with recent preliminary observations in humans showing that atorvastatin might ameliorate nocturnal penile erection (Saltzman et al, 2004) and sildenafil responsiveness (Herrmann et al, 2006) in hypercholesterolemic patients with ED. Whether atorvastatin potentiation of sildenafil-induced erections in SHR is due to the above-described effects of the statin on the RhoA/ROCK pathway or involves positive effects on endothelial function and NO signaling (Bonetti et al, 2003; Rikitake et al, 2005) needs to be established.
It is interesting to note that elocalcitol, a vitamin D analog chemically unrelated to statins but known to negatively interfere with RhoA/ROCK in the bladder (Morelli et al, 2007), was unable to restore sildenafil responsiveness in SHR. In addition, in isolated penile SM cells, elocalcitol inhibited cell migration and proliferation at concentrations 3 log units higher than in other SM cells, such as those derived from prostate (Crescioli et al, 2003a) and bladder (Crescioli et al, 2005). A possible explanation for this lower sensitivity of penile musculature to elocalcitol, compared with other urinary tissues, is provided by the finding that in SHR, VDR expression is more abundant in the bladder than in the penis. Because elocalcitol actions are VDR mediated, this might explain why elocalcitol ameliorates bladder overactivity in SHR (Morelli et al, 2007) but not altered penile erections, as shown in the present study. Thus, the differential quantitative expression of VDR in bladder and CC suggests a plausible mechanism for the tissue-specific effect of elocalcitol on the RhoA/ROCK contractile pathway. Interestingly, elocalcitol has been reported to possess tissue- and cell-type selectivity in VDR activation, acting as a poor VDR agonist in tissues like intestine and kidney but as a potent VDR agonist in bone (Peleg et al, 2002, 2003). Our findings further extend the tissue selectivity of elocalcitol and show its ability to modulate RhoA/ROCK signaling only in defined target tissues. These cell context–selective actions, reported also for other VDR agonists (Ma et al, 2006), translate into reduced induction of hypercalcemia and may account for the placebo-like side effect profile induced by elocalcitol treatment in patients (Colli et al, 2006).
In conclusion, our data indicate that atorvastatin, but not elocalcitol, ameliorates sildenafil-induced penile erections in SHR, most probably by interfering with RhoA/ROCK signaling. This pleiotropic activity of atorvastatin, independent from the lipid-lowering one, might explain its positive effects in restoring erectile function and sildenafil responsiveness in ED patients (Saltzman et al, 2004; Herrmann et al, 2006). Accordingly, a recent report indicates that atorvastatin can improve postoperative erectile function even in non-hypercholesterolemic patients undergoing radical retropubic prostatectomy (Hong et al, 2007).
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Footnotes |
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|| These authors contributed equally to the article. 
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References |
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7
rats/group. Closed bars indicate WKY; open bars, untreated spontaneously
hypertensive rats (SHR); diagonally hatched bars, atorvastatin-treated SHR (5
mg/kg for 2 weeks); horizontally hatched bars, atorvastatin-treated SHR (30
mg/kg for 2 weeks); grey bars, elocalcitol-treated SHR (30 µg/kg for 2
weeks). °P < .05 and °°P < .01 vs WKY
rats. *P < .05 vs untreated SHR.
