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. Author manuscript; available in PMC: 2010 Mar 2.
Published in final edited form as: Biol Reprod. 2006 Sep 20;76(1):19–28. doi: 10.1095/biolreprod.106.053926

Regulation of Cavernous Nerve Injury-Induced Apoptosis by Sonic Hedgehog1

Carol A Podlasek 3,2, Cynthia L Meroz 4, Yi Tang 4, Kevin E McKenna 3,4, Kevin T McVary 3
PMCID: PMC2830895  NIHMSID: NIHMS175892  PMID: 16988214

Abstract

Thirty to eighty-seven percent of patients treated by radical prostatectomy experience erectile dysfunction (ED). The reduced efficacy of treatments in this population makes novel therapeutic approaches to treat ED essential. We propose that abundant apoptosis observed in penile smooth muscle when the cavernous nerve (CN) is cut (mimicking the neural injury which can result from prostatectomy) is a major contributing factor to ED development. We hypothesize that decreased Sonic hedgehog (SHH) signaling is a cause of ED in neurological models of impotence by increasing apoptosis in penile smooth muscle. We examined this hypothesis in a bilateral CN injury model of ED. We found that the active form of SHH protein was significantly decreased 1.2-fold following CN injury, that SHH inhibition causes a 12-fold increase in smooth muscle apoptosis in the penis, and that SHH treatment at the time of CN injury was able to decrease CN injury-induced apoptosis (1–3-fold) in a dose-dependent manner. These results show that SHH stabilizes the alterations of the corpora cavernosal smooth muscle following nerve injury.

Keywords: apoptosis, male sexual function, penis, signal transduction

INTRODUCTION

Erectile dysfunction (ED) is a serious medical condition that affects 52% of men between the ages of 40 and 70. The incidence of ED increases with age, coronary artery disease, peripheral vascular disease, smoking, dyslipidemia, higher BMI, diabetes mellitus, and postradical prostatectomy [1]. Forty to seventy percent of prostate cancer patients treated by radiotherapy [2, 3] and 30%–87% of patients treated by radical prostatectomy experience ED [3–6]. Although potency improves with time postprostatectomy [7], sexual dysfunction was common 5 years following radical prostatectomy [8]. Current treatment options for ED, including self-injection or intra-urethral administration of alprostadil, vacuum erection devices, and phosphodiesterase 5a (PDE5A) therapy, have proven to be only partially effective [6, 9–10]; PDE5A inhibitors are ineffective in 29%–86% of prostatectomy patients who experience ED, depending on their nerve injury status [11]. The reduced efficacy of all treatments makes prostatectomy patients a population for whom novel therapeutic approaches to treat ED are needed. Smooth muscle cell atrophy is abundant in PDE5A nonresponders [12]. We propose that extensive morphological changes in penile smooth muscle and endothelium observed postprostatectomy in the corpora cavernosa are a major contributing factor to ED development. Interventions targeted to prevent cavernous nerve (CN)-induced apoptosis in the penis would be a powerful tool to prevent postprostatectomy ED.

The process of erection is complex, involving critical integration of vascular, neural, hormonal, and morphological influences. As ED develops, the balance between these processes becomes skewed. In both diabetic and CN injury ED models, there is a general impairment in responsiveness of the rat corpora cavernosa [13], impaired smooth muscle relaxation [14], a significant decrease in abundance of the smooth muscle and endothelium of the corpora cavernosa [15, 16], and an increase in the ratio of apoptotic cells in the erectile tissue [16–18]. Current treatments for ED aim to increase smooth muscle relaxation. However, as the smooth muscle morphology of the corpora cavernosa becomes increasingly abnormal following prostatectomy, these traditional treatment strategies become less effective and eventually fail. In these studies we propose a novel approach, in which we aim to elucidate the underlying mechanisms that cause apoptosis of the corpora cavernosal smooth muscle, and thus cause ED to occur. If penile apoptosis could be prevented following prostatectomy while the CN regenerates, then resumption of normal erectile function would occur more quickly and fibrosis would be prevented as the tissue tries to regenerate. This would lead to long-term prevention of ED. Investigation of the factors that regulate apoptosis in the penis and the role that neural innervation plays in this process is a necessity to move the field of ED research forward and to develop new treatment strategies. A recently identified regulator of penile morphology [19] that has been shown to orchestrate apoptosis induction during embryogenesis of the penis [20, 21] is the morphogen Sonic hedgehog (SHH).

We have shown in previous studies that the SHH signaling pathway is critical for establishing the sinusoid morphology of the corpora cavernosa during development [19] and that SHH continues to function in the adult organ to regulate and maintain penile morphology [16, 19]. Inhibition of SHH signaling in normal adult rats caused abundant apoptosis and remodeling of the sinusoidal architecture of the corpora cavernosa, so that sinusoids were completely absent [19]. The morphological changes after SHH inhibition were so extensive that they affected penile physiology and caused significant ED [19]. We propose the hypothesis that decreased SHH signaling is a cause of ED in neurological models of impotence by increasing apoptosis in penile smooth muscle, which leads to morphological changes within the corpora cavernosal sinusoidal tissue, and thus to ED. This hypothesis is supported by previous observations of decreased SHH protein, smooth muscle and endothelium, significantly increased apoptosis, and erectile dysfunction in BB/WOR diabetic rats [16, 22]. In this study we have examined a second model of ED, the bilateral CN-injured Sprague Dawley rat, in order to determine if decreased SHH signaling derived from the nerves is the cause of morphology changes (increased apoptosis, smooth muscle and endothelial degradation) which occur in the corpora cavernosa postprostatectomy, and to evaluate if SHH protein treatment of the penis can be used to prevent CN injury-induced apoptosis. In this study we have shown that the active form of SHH protein was significantly decreased following CN injury and that SHH treatment at the time of CN injury was able to prevent/postpone CN injury-induced apoptosis. These results show that SHH has significant potential to be developed as a treatment to prevent smooth muscle apoptosis in the penis postprostatectomy.

MATERIALS AND METHODS

Animals

Sprague-Dawley rats, postnatal day 120 (P120), were obtained from Charles River. Animals were cared for in accordance with the National Research Council publication Guide for Care and Use of Laboratory Animals.

Cavernous Nerve Injury

P120 Sprague-Dawley rats were randomized into two groups: bilateral CN resection (n = 46) and sham abdominal exploration (control, n = 46). Five-millimeter sections of the cavernous nerve were removed bilaterally using a KAPS industrial microscope under direct vision through a midline abdominal incision. The prostatic capsule was manipulated in control animals without resecting the cavernous nerve. Stress-related fluctuations of serum testosterone were minimized at the time of abdominal exploration through bilateral epididymo-orchiectomy and subcutaneous placement of a 2-cm piece of medical grade Silastic tubing (Dow Corning Corp., Midland, MI) filled with crystalline testosterone [23, 24]. This method ensures reliable, uniform serum testosterone levels for both the control and intervention groups up to 28 days after placement [25]. Penes were harvested from euthanized males by sharp dissection 2, 4, 7, 14, and 21 days after CN resection and were either frozen in liquid nitrogen or fixed in 4% paraformaldehyde.

Western Analysis of Control and CN-injured Corpora Cavernosa

Western analysis assaying for SHH protein was performed on protein samples isolated from corpora cavernosa from control (n = 5) and CN-injured (n = 5) Sprague Dawley rat penes 21 days after CN injury, as outlined previously [16, 19]. Proteins were separated via electrophoresis using a 10% polyacrylamide gel and transferred to a nitrocellulose membrane (Bio-Rad) using a Hoefer SemiPhor Semi-Dry Transfer Unit (Amersham Pharmacia, Piscataway, NJ) for 1 h. Membranes were blocked for 1 h at room temperature in 5% nonfat skim milk in PBS Tween buffer. Membranes were incubated with either 1:50 SHH (N-19, SC-1194, Santa Cruz) or 1:50 000 ACTB (formerly known as β-ACTIN, Sigma) antibodies overnight at 4°C. Membranes were washed with PBS-Tween one time for 10 min and then incubated with 1:45 000 anti-goat and 1:70 000 anti-mouse secondary antibodies for 1 h at room temperature. Protein bands were visualized using HRP-conjugated anti-biotin (ECL western blotting detection reagent, Amersham) according to manufacturer’s directions and were exposed to Hyperfilm. Bands were quantified by densitometry using Kodak 1D software (Rochester, NY). Quantification of bands was performed by determining the ratio of the density of SHH divided by ACTB to eliminate differences in protein loading. Significant differences in protein abundance of SHH/ACTB were determined in control and CN-injured penes using a t-test (Microsoft’s Excel program). Samples were run five times, and the ratios for each sample were averaged.

Immunohistochemical Analysis

Immunohistochemical analysis (IHC) was performed as previously outlined [16, 19] on adult Sprague Dawley rats (P120, n = 6) assayed for goat polyclonal SHH (N-19, SC-1194, Santa Cruz), SHH (C-18, SC-1195, Santa Cruz), and PTCH1 (G-19, SC-6149, Santa Cruz) using a 1/100 dilution of primary antibody. Dual staining was performed using SHH (N-19)/PECAM1 (formerly known as CD31, mouse monoclonal antibody, Chemicon International), SHH (N-19)/ACTA1 (formerly known as α-ACTIN, mouse monoclonal antibody, Sigma-Aldrich), SHH(N-19)/prolyl 4-hydroxylase (P4HB, mouse antibody, ACRIS Antibodies), PTCH1 (G-19)/PECAM1 (mouse monoclonal antibody, Chemicon International), and PTCH1 (G-19)/ACTA1 (mouse monoclonal antibody, Sigma-Aldrich) using a 1/100 dilution of primary antibodies. Secondary antibodies used were Alexa Fluor 488 rabbit anti-goat (1/300, Molecular Probes) and Alexa Fluor 594 chicken anti-mouse (1/300, Molecular Probes). Negative controls were performed with secondary only (without primary) to test for nonspecific staining and autofluorescence. Sections were mounted using Pro-Tex Mounting Medium (Baxter Diagnostics, Inc., Pittsburgh, PA). Microscopy was performed using a dual light and fluorescent microscope (Leitz) and photographed using a Nikon digital camera.

Control (n = 6) and CN-injured (n = 6) penes 21 days after nerve injury were assayed with SHH (N-19, Santa Cruz) and PTCH1 (G-19, Santa Cruz) as described above.

TUNEL Assay for Apoptosis

Staining for apoptosis was performed on control (n = 4 at each time point) and CN-injured (n = 4 at each time point) penes 2, 4, 7, and 21 days after CN injury (CN21, respectively), using the ApopTag kit (Intergen, Purchase, NY). Fluorescent apoptotic cells were observed under a fluorescent microscope (Leitz) and photographed using a Nikon digital camera.

Electron Microscopy

Electron microscopy was performed as described previously [26]. Control (n = 4) and CN-injured (n = 4) penes 21 days after CN injury were fixed in 2.5% glutaraldehyde, postfixed in 1% OsO4, dehydrated, and embedded in Epon resin. Thin sections were cut and stained with 2% uranyl acetate and 3% lead citrate. Electron microscopy was performed using a JEOL 100CX Transmission Electron Microscope to identify in which cell type apoptosis was taking place. Apoptosis was identified by the presence of condensed chromatin, nuclear fragmentation, and cytoplasmic blebbing, which are common in cells undergoing apoptosis [27].

Reversibility of SHH Inhibition

Affi-Gel beads (100–200 mesh, Bio-Rad Laboratories, Hercules, CA) were equilibrated with 5E1 SHH inhibitor (n = 3 at each time postinjection, 1–3 µg/ml, Jessel, Hybridoma bank at the University of Iowa), or PBS (control, n = 2 at each time postinjection) overnight at 4°C. Approximately 30–40 beads were injected directly into the corpora cavernosa of P120 Sprague Dawley rat penes. Rats used for these experiments were normal rats that had not undergone CN injury. Rats were killed 10 days, 4 wk, and 6 wk following Affi-gel bead injection (0, 2, and 4 wk after SHH inhibitor in the beads was depleted). Four weeks after bead injection was chosen as a time point to examine if SHH inhibition was reversible, since it represented a comparable amount of time for the tissue to recover as it was exposed to SHH inhibitor. This was insufficient time for complete recovery, so an additional time point (6 wk after bead injection) was added to ensure complete recovery to normal morphology after SHH inhibition. Penes were harvested, fixed in 4% paraformaldehyde, and embedded in paraffin for sectioning. IHC was performed on control and SHH inhibitor-treated penes assaying for SHH (N-19, SC-1194, Santa Cruz) and ACTA1 (Sigma, St. Louis, MO) proteins as described above. Bead technology has previously been used successfully for delivery of proteins and antibodies to target tissues [28, 19].

SHH Protein Injection at the Time of CN Injury

Affi-Gel beads (100–200 mesh, Bio-Rad Laboratories, Hercules, CA) were equilibrated with either 1× recombinant mouse SHH peptide [29] (n = 10, 7.5 µg per animal, R & D Systems), 2× SHH peptide (n = 9, 15 µg per animal, R & D Systems), or PBS (control, n = 9) overnight at 4°C. Approximately 30–40 beads were injected directly into the corpora cavernosa of control and SHH peptide-treated P120 rat penes at the time of bilateral CN injury surgery (described above). Rats were killed 2, 4, and 8 days following surgery and bead injection. CN injury only (positive control, n = 10) rats served as a positive control for apoptosis following CN injury. Additional controls were performed as follows: heat-inactivated 1× (n = 3) and 2× (n = 3) SHH protein killed at 4 and 8 days post-CN. All penes were harvested and fixed in 4% paraformaldehyde for sectioning. TUNEL staining for apoptosis was performed as described above. Quantification of apoptosis was performed by counting the number of apoptotic cells in photographs of 3–5 fields per rat. All cells in a given field were counted in a similar manner based on propidium iodide staining. Changes in the number of apoptotic cells were reported as the ratio of the average number of apoptotic cells to the average number of all cells. Significance was determined using a t-test. The ratio of apoptotic cells to all cells was difficult to quantify reliably, since it was dependent on the distance from Affi-Gel beads, the number of beads in a given area, and the amount of SHH protein delivered to a particular area of the penis. An attempt was made to alleviate these concerns as much as possible by quantifying apoptosis within a given distance relative to the Affi-Gel beads (330 µm).

Effect of SHH Inhibition and Supplementation on VEGFA Signaling in Intact Rats

Affi-Gel beads (100–200 mesh, Bio-Rad Laboratories, Hercules, CA) were equilibrated with either 5E1 SHH inhibitor (n = 5, 1–3 µg/ml, Jessel, Hybridoma bank at the University of Iowa), recombinant mouse SHH peptide [29] (n = 5, 7.5 µg per animal, R & D Systems), or mouse IgG (control, n = 5, 3 µg/ml) overnight at 4°C. Approximately 30–40 beads were injected directly into the corpora cavernosa of P120 rat penes. Sprague Dawley rats used for these experiments were normal rats that had not undergone CN injury. Rats were killed 7 days postinjection. Penes were harvested and fixed in 4% paraformaldehyde for sectioning. IHC was performed on control, SHH protein, and SHH inhibitor-treated penes assaying for vascular endothelial growth factor (VEGFA, goat polyclonal antibody, Santa Cruz, 200 µg/ml and mouse monoclonal antibody, Chemicon International) as described above. Sections were stained with diaminobenzidine (DAB) and mounted using Pro-Tex Mounting Medium (Baxter Diagnostics, Inc., Pittsburgh, PA). TUNEL staining for apoptosis was also performed on control, SHH-inhibited, and SHH protein-treated penes as described above. Quantification of apoptosis was performed by counting the number of apoptotic cells in photographs of 3–5 fields per rat and was reported as the average number of apoptotic cells. Significance was determined using a t-test. The number of apoptotic cells was difficult to quantify reliably, since it was dependent on the distance from Affi-Gel beads, the number of beads in a given area, and the amount of SHH protein or SHH inhibitor delivered to a particular area of the penis. An attempt was made to alleviate these concerns as much as possible by quantifying apoptosis within a given distance relative to the Affi-Gel beads (130 µm).

RNA Isolation and Quantification of Gene Expression by RT-PCR

Total RNA was extracted from individual intact penes of CN-injured and control Sprague Dawley rats using the TRIzol (Life Technologies, Gaithersburg, MD) method. Samples were DNAse (Promega)-treated to eliminate genomic DNA contamination. Primers [16] were purchased from Integrated DNA Technologies, Inc. (Coralville, IA). RT-PCR was performed using the GeneAmp RNA PCR Core Kit (Perkin-Elmer, Branchburg, NJ), and products were restriction-digested to confirm that bands represented the sequences of interest. Quantitative RT-PCR was performed as described previously [30, 16, 19] using noncompetitive methodology and glyceraldehyde-3-phosphate dehydrogenase (Gapdh) as an endogenous internal standard. All measurements were made in the linear range for Shh and Gapdh (n = 9 control and 9 CN-injured penes) and Ptch1 and Gapdh (n = 4 control and n = 4 CN-injured penes). Bmp4 expression was also measured (n = 7 control and n = 8 CN-injured penes) in the linear range for Bmp4 and ribosomal protection subunit 32 (Rpl32), which was used as an endogenous internal standard. Assays were performed in triplicate on individual tissue specimens, the results averaged, and the product ratios reported as the mean plus or minus SEM. In order to compare Shh, Ptch1, and Bmp4 expression in the 21-day control and CN-injured rats, the data were normalized to 1 (maximum value set equal to 1). Shh expression was also measured at 7 (n = 6 control and 6 CN-injured) and 14 days (n = 8 control and 8 CN-injured) after CN injury. Normalization to 1 was not performed, since data were collected in one experiment. Excel (Microsoft) was used for statistical analysis, and a t-test was used to determine significant changes in RNA expression.

In Situ Hybridization

In situ hybridization was performed as previously described [31, 32, 16, 19] on control (n = 6) and CN-injured (n = 6) penes 21 days after CN injury. Penes were fixed in 4% paraformaldehyde overnight. A mouse Shh RNA probe was obtained from Andrew McMahon [29] and a mouse Ptch1 RNA probe from Matthew Scott [33].

Statistical Analysis

All results are expressed as the average ± SEM. Differences were analyzed by t-test and considered statistically significant at P < 0.05.

RESULTS

Localization of SHH and PTCH1 in the Adult Penis by IHC

Antibodies that recognize the active and precursor forms of SHH protein and PTCH1 were used to determine their localization in adult Sprague Dawley penis tissue. N-terminal SHH protein (active and precursor forms) was localized within the corpora cavernosal sinusoidal tissue, a layer adjacent to the tunica, Schwann cells of the nerves, veins (Fig. 1), and the urethra [19]. Colocalization of SHH and ACTA1 by dual IHC identifies SHH in smooth muscle cells of the corpora cavernosal sinusoidal tissue. SHH is present to a lesser extent in cells residing between the sinusoids of the corpora. These cells which stain for SHH protein colocalize with P4HB, a fibroblast marker (Fig. 2). The localization of PTCH1 protein was very similar to SHH (Fig. 1 and Fig. 2), with staining present within the corpora cavernosal sinusoidal tissue, a layer adjacent to the tunica, Schwann cells of the nerves, veins and the urethra [19]. Colocalization of PTCH1 and ACTA1 by dual IHC identify PTCH1 in smooth muscle cells of the corpora cavernosal sinusoidal tissue. The localization of the C-terminal SHH protein (precursor form only, Fig. 1) was similar to N-terminal SHH, with two notable differences. Precursor SHH protein was abundant in the perineurium of the nerves, but was not present in the Schwann cells or the veins, as was the active form of SHH protein. SHH/PECAM1 and PTCH1/PECAM1 dual staining shows that SHH and PTCH1 do not colocalize with PECAM1, indicating that SHH and PTCH1 are not present in endothelial cells.

FIG. 1.

FIG. 1

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IHC analysis of SHH (N-19 and C-18) and PTCH1 in adult Sprague Dawley rat penes. a) The precursor and active forms of SHH protein recognized by SHH (N-19) are abundant within corpus cavernosal sinusoidal tissue and a layer under the tunica (left), nerves of the dorsal nerve bundle, and veins (middle). n, Nerves; v, vein. b) The localization of PTCH1 is similar to the active form of SHH. c) The localization of the precursor SHH protein (C-18) is similar to that of the active form. However, it is not present in the veins and is restricted to the perineurium of the nerves. Elastin fibers in the veins autofluoresce and do not stain for SHH protein. Original magnification ×60.

FIG. 2.

FIG. 2

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Dual IHC for SHH/ACTA1, SHH/PECAM1, SHH/P4HB, PTCH1/ACTA1, and PTCH1/PECAM1 in adult Sprague Dawley rat penes. SHH and PTCH1 proteins colocalize with ACTA1 in the corpus cavernosal sinusoidal tissue. In addition, SHH protein is present in cells located between the sinusoids. These cells stain for P4HB, a fibroblast marker. SHH and PTCH1 proteins are shown in green, and ACTA1, PECAM1, and P4HB proteins are presented in red. Yellow represents dual staining. Neither SHH nor PTCH1 colocalize with PECAM1, indicating that SHH and PTCH1 are not present in endothelial cells. Original magnification ×400 (ACTA1 and PECAM1) and ×1000 (P4HB).

Shh and Ptch1 RNA Expression Increased Significantly by RT-PCR Analysis Following CN Injury

Relative changes in RNA expression of Shh and Ptch1 were quantified by RT-PCR in control and CN-injured penes (Fig. 3). Shh and Ptch1 RNA expression significantly increased 1.25-fold and 2-fold, respectively, 21 days after CN injury (P-values = 0.01 and 0.002, respectively). Shh RNA expression remained unchanged when assayed at 7 (control = 0.25 ± 0.05, CN-injured = 0.39 ± 0.1, P-value = 0.13) and 14 days post-CN injury (control = 0.46 ± 0.1, CN-injured = 0.32 ± 0.08, P-value = 0.12). Bmp4 expression was also measured 21 days after CN injury. Bmp4 expression was unchanged following CN injury (control = 0.62 ± 0.08, CN-injured = 0.71 ± 0.09, P-value = 0.25).

FIG. 3.

FIG. 3

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Shh and Ptch1 RNA expression in control and CN-injured rats. Relative abundance of Shh and Ptch1 RNA expression measured by semiquantitative RT-PCR analysis in control and CN-injured Sprague Dawley rat penes 21 days after CN injury. Shh and Ptch1 expression were significantly increased 21 days after CN injury (P-value = 0.01 and 0.002, respectively). Asterisks identify significant changes in expression. C, Control; CN, cavernous nerve injury.

Localization of Shh and Ptch1 RNA by In Situ Hybridization Remained Unchanged Following CN Injury

The RNA localization of Shh and Ptch1 was examined by in situ hybridization in control and CN-injured penes 21 days after CN injury. Shh and Ptch1 RNA expression was abundant in the smooth muscle of the corpora cavernosal sinusoidal tissue of both control and CN-injured penes (Fig. 4). A change in RNA localization for Shh and Ptch1 was not observed following CN injury.

FIG. 4.

FIG. 4

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In situ hybridization showing the localization of Shh and Ptch1 in control and CN-injured Sprague Dawley rat penes 21 days after CN injury. The distribution of Shh and Ptch1 RNA remained unchanged following CN injury in smooth muscle cells of the corpus cavernosal sinusoidal tissue. Original magnification ×400. Arrows indicate Shh and Ptch1 RNA localization.

The Active Form of SHH Protein was Significantly Decreased by Western Analysis Following CN Injury

The precursor (46 kDa) and active (19 kDa) forms of SHH protein were quantified by Western analysis in control and CN-injured penes 21 days after CN injury. The active form of SHH protein was significantly decreased after CN injury (control = 0.572 ± 0.02, CN-injured = 0.492 ± 0.01, P-value = 0.001, Fig. 5). The precursor form was increased, but there was not enough statistical power for the difference to be significant (control = 0.815 ± 0.03, CN-injured = 0.879 ± 0.03, P-value = 0.08, Fig. 5). An additional band at 38 kDa was observed that was blocked using the antigenic peptide for SHH. From the size of the product it is hypothesized that this band represents a multimer of the active SHH protein. SHH multimerization has been reported in the literature [34]. The 38-kDa product was unchanged with CN injury (control = 1.070 ± 0.04, CN-injured = 1.070 ± 0.03, P-value = 0.5, Fig. 5). The localization of SHH and PTCH1 remain unchanged after CN injury by IHC analysis.

FIG. 5.

FIG. 5

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Graph (a) and gel (b) showingWestern analysis of SHH protein in control (C) and CN-injured (CN) penes 21 days after CN injury. The active form of SHH protein (19 kDa) was significantly decreased after CN injury (P-value = 0.001). The precursor form (46 kDa) was increased but was not statistically significant (P-value = 0.08). SEM for the 19-kDa bands was 0.02, which was too small to print error bars on the graph.

Morphological Changes in the Corpora Cavernosa Caused by SHH Inhibition Are Reversible

The corpora cavernosa of adult Sprague Dawley rats were treated with 5E1 SHH inhibitor by Affi-Gel bead injection into the corpora cavernosa. Ten days after injection, the corpora cavernosa had remodeled so that sinusoids were absent in the treated region (Fig. 6). SHH inhibitor was depleted from the beads after 10–14 days. At 4 and ± wk postinjection, penile morphology was examined and SHH and ACTA1 proteins were assayed by IHC. After 4 wk, sinusoidal morphology was partially reestablished and SHH and ACTA1 were observed in the corpora cavernosa (Fig. 6). After ± wk, smooth muscle lined sinusoidal tissue was abundant, the morphology of the corpora cavernosa was indistinguishable from PBS-treated controls (Fig. 6), and SHH protein was present in the sinusoidal tissue (data not shown).

FIG. 6.

FIG. 6

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IHC analysis of ACTA1 staining for smooth muscle in SHH inhibited Sprague Dawley penes 10 days, 4 wk, and 6 wk after injection of SHH inhibitor and PBS, which was used as a control for the bead vehicle. a) The absence of sinusoids and positive staining for smooth muscle are evident during SHH inhibition (1–14 days) in the corpora cavernosa. b) Four weeks after SHH inhibitor injection, the presence of sinusoidal tissue staining positively for ACTA1 was identified. c) Six weeks after SHH inhibitor injection, the morphology of the sinusoidal tissue closely resembled normal corpora cavernosa morphology and was indistinguishable from the PBS-treated control corpora cavernosa (d). Original magnification ×60.

CN Injury Induces Abundant Apoptosis

TUNEL staining was performed on control and CN-injured penes 2, 4, 7, and 21 days following CN injury in Sprague Dawley rat penes. Apoptosis was abundant the first week following CN injury (Fig. 7) and remained elevated above basal levels at 21 days post-CN. Electron microscopy identified abundant apoptosis in smooth muscle cells and to a lesser extent in endothelial cells of the corpora cavernosa following CN injury (Fig. 7).

FIG. 7.

FIG. 7

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TUNEL staining of CN-injured, SHH inhibited, and SHH protein-treated/CN-injured rats. a) TUNEL staining of corpus cavernosal sinusoidal tissue from CN-injured penes 4 days post-CN injury (left). Apoptosis is abundant in and around the sinusoidal tissue. White arrows indicate apoptotic cells. Original magnification ×400. Electron microscopy was used to examine apoptosis in CN-injured Sprague Dawley rat penes 21 days after CN injury (middle and right). Apoptosis was identifiable in endothelial cells (middle, original magnification ×4400) and abundant in smooth muscle cells (right, original magnification ×4400) of the corpora cavernosa. Apoptotic cells were identified by the appearance of condensed chromatin, membrane blebbing, and nuclear degradation. sm, Smooth muscle; e, endothelium; s, sinusoidal space. Arrows indicate apoptotic cells. b) TUNEL staining of SHH-inhibited (left) and PBS-treated control (right) corpora cavernosa from adult Sprague Dawley rat penes. Apoptosis was abundant following SHH inhibition in the corpora cavernosa, and the localization of apoptosis was very similar to that observed following CN injury. Apoptosis was not evident in PBS-treated control penes, indicating that the presence of the Affi-Gel bead vehicles used to deliver SHH inhibitor or PBS to the corpora cavernosa did not induce apoptosis. Original magnification ×400. White arrow indicates apoptotic cells, and yellow arrows indicate red blood cells, which autofluoresce. c) TUNEL staining of Sprague Dawley corpora cavernosa that were treated with PBS (control, left) or SHH protein (right) via Affi-Gel bead vehicle at the time of bilateral CN injury. Four days following CN injury, apoptosis was abundant in the corpora cavernosa, as observed in the PBS-treated control penes (left). This shows that the presence of the bead vehicle itself did not decrease the amount of apoptosis induced by CN injury. In the SHH protein-treated corpora cavernosa, the presence of apoptotic cells was significantly decreased following CN injury (right). White arrows indicate apoptotic cells, and yellow arrows indicate red blood cells, which autofluoresce. b, Affi-Gel bead. Original magnification ×100; bar = 10 µm.

SHH Inhibition in the Corpora Cavernosa Causes Apoptosis

TUNEL staining for apoptosis was performed on penes from adult Sprague Dawley rats that had been treated with SHH inhibitor for 7 days via Affi-Gel beads or control rat penes treated with PBS. Apoptosis was abundant within the sinusoidal tissue and the tissue between the sinusoids of the corpora cavernosa following SHH inhibition (Fig. 7, number of apoptotic cells within 130 µm of a bead in 5E1 SHH inhibitor= 44.3 ± 5.2 and in PBS control = 3.8 ± 0.8 treated penes, P-value = 0.0001). Even sinusoidal tissue distant from the Affi-Gel bead vehicles showed abundant apoptosis after SHH inhibitor treatment. Apoptosis was not induced in control rats (Fig. 7) or in SHH protein-treated rats (SHH protein-treated = 1.6 ± 0.7).

SHH Protein Treatment at the Time of CN Injury Prevents Post-CN Injury-induced Apoptosis

Affi-Gel beads soaked in SHH protein were injected into the corpora cavernosa at the time of bilateral CN injury in adult Sprague Dawley rats. After 2 days, apoptosis was abundant and there was no difference in the number of apoptotic cells in the presence or absence of SHH protein (ratio of apoptotic cells to all cells within ~330 µm of a bead in SHH protein CN2 = 0.85 ± 0.11 and in PBS CN2 = 0.82 ± 0.06 treated penes). At 4 days post-CN injury, the amount of apoptosis was decreased in the presence of SHH protein (Fig. 7, ratio of apoptotic cells to all cells within ~330 µm of a bead in SHH protein-treated CN4 = 0.64 ± 0.05 and PBS control = 0.8 ± 0.06 penes, P-value = 0.02). When double the concentration of SHH protein was applied, apoptosis was suppressed in a larger region surrounding the bead vehicles and apoptosis was further reduced by 2.5-fold (ratio of apoptotic cells to all cells within ~330 µm of a bead in 2× SHH protein-treated CN4 = 0.33 ± 0.09 and control heat-inactivated 2× SHH protein-treated CN4 = 0.82 ± 0.03 penes, P-value = 3.39E-05). At 8 days post-CN injury, apoptosis was still suppressed in the presence of SHH protein (ratio of apoptotic cells to all cells within ~330 µm of SHH protein-treated CN8 = 0.24 ± 0.07 and control heat-inactivated SHH protein-treated CN8 = 0.71 ± 0.06 penes, P-value = 9.03E-05). CN injury-induced apoptosis was not suppressed in control penes. The amount of apoptosis suppression was dependent on the distance from the Affi-Gel beads at ;95 µm (ratio of apoptotic cells to all cells for control heat-inactivated 2× SHH protein CN4 = 0.80 ± 0.03 and 2× SHH protein CN4 = 0.05 ± 0.03, P-value = 4.32E-05), ~215 µm (ratio of apoptotic cells to all cells for control heat-inactivated 2× SHH protein CN4 = 0.78 ± 0.05 and 2× SHH protein CN4 = 0.38 ± 0.14, P-value = 0.02), and at ~330 µm (ratio of apoptotic cells to all cells for control heat-inactivated 2× SHH protein CN4 = 0.91 ± 0.04 and 2× SHH protein CN4 = 0.57 ± 0.07, P-value = 0.008) from the beads.

SHH Induction of VEGFA

Adult Sprague Dawley rat penes were treated with either 5E1 SHH inhibitor or SHH protein via Affi-Gel beads, and IHC was performed on penis tissue assaying for VEGFA. VEGFA protein was upregulated in response to SHH protein treatment and was reduced in the presence of SHH inhibitor (Fig. 8).

FIG. 8.

FIG. 8

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IHC analysis of VEGFA localization in control (top), SHH-protein-treated (middle), and SHH-inhibited (bottom) adult Sprague Dawley rat penes. VEGFA protein is normally abundant within the corpus cavernosal sinusoidal tissue (original magnification ×400). VEGFA protein was upregulated after SHH protein treatment and downregulated after SHH inhibition in the corpora cavernosa (original magnification ×250), suggesting that SHH may be an upstream regulator of VEGFA. Arrows indicate VEGFA protein; b, Affi-Gel bead.

DISCUSSION

Genes involved in cell fate decisions during development also play key roles in continuous cell fate decisions made by adult organs. The Shh signaling pathway is critical for establishing the sinusoid morphology of the corpora cavernosa, and Shh continues to regulate and maintain penile morphology in the adult organ [19]. In two rat models of ED, the BB/WOR diabetic rat and in the CN-injured Sprague Dawley rat, SHH protein is significantly decreased [16]. In these same models there are significant morphological changes in the corpora cavernosa, including increased apoptosis and decreased smooth muscle and endothelial staining [19, 32, 18]. These changes are identical to those observed after SHH inhibition using a chemical inhibitor in normal rats. If decreased SHH protein is a cause of decreased smooth muscle and ED in diabetic and CN-injured penes, then examining the potential reversibility of SHH inhibition is of critical clinical significance. The experiments in this study provide evidence that the morphological changes caused by SHH inhibition are reversible in the rat with time, suggesting that the accompanying ED may also be reversible with reestablished/supplemented SHH signaling. This shows that SHH has significant potential for development as a therapy for morphological and physiological changes in the penis that cause ED.

The reversibility of SHH inhibition has wide implications. Mutations in the Shh signaling pathway that target Ptch1, Smo, and Gli1 are associated with certain forms of cancer, including prostate, skin, and esophageal [35–37]. These mutations cause continuous transcription of Shh targets and are not caused by an overabundance of Shh itself. The SHH inhibitor cyclopamine is currently being tested to fight tumor growth. If given systemically, this inhibitor will likely alter normal penile morphology and the architecture of other organs where Shh remains active in the adult. Penile remodeling occurs quickly after SHH inhibition and is extensive after 7 days. The morphological changes caused by SHH inhibition in the penis are reversible, at least after short-term inhibition (< 14 days). This suggests that SHH inhibitors given to prostate cancer patients will modify normal penile morphology and cause ED. However this process may be reversible once SHH inhibition is removed.

It is an interesting finding that the RNA expression and precursor form of SHH protein increase while the active form of SHH protein decreases following CN injury. This suggests that there is inhibited posttranslational processing of the SHH protein with nerve injury and feedback regulation on Shh RNA expression in order to replace the decreased protein. Feedback regulation of Shh by downstream targets, including Bmp4 and Ptc, have been reported in other systems in the literature [38–40]. Alteration in processing may occur at several levels, including the autocatalytic cleavage of the precursor to form the cholesterol-modified 19-kDa active SHH product; the multimerization of SHH, which localizes the protein to the lipid rafts of the cell membrane; and the release of the multimeric form from the membrane, which effects long-range SHH signaling. While beyond the scope of this study, identification of the mechanism by which the active form of SHH protein is decreased following CN injury is an interesting and important avenue for further research, since it will shed light on how altered SHH signal transduction may lead to abnormal penile morphology.

In this work, we continue our ongoing efforts to determine what role neuropathy plays in ED development and to better understand the molecular mechanisms that regulate maintenance of penile architecture. We have shown that SHH inhibition causes apoptosis and that SHH protein is significantly decreased following CN injury. The localization of apoptosis in the corpora cavernosa after CN injury and after SHH inhibition appears very similar, with apoptotic cells present in the sinusoid smooth muscle and endothelium (Fig. 7). These results suggest that decreased SHH protein may be a cause of the abundant apoptosis that occurs following CN injury. They also suggest that corpora cavernosa SHH signaling is regulated by neural innervation. When SHH protein was introduced into the corpora cavernosa at the time of CN injury, SHH was able to prevent apoptosis in a dose-dependent manner. These results show that SHH stabilizes the alterations of the corpora cavernosal smooth muscle following nerve injury. If penile apoptosis could be prevented while the cavernous nerve regenerates, the potential for preservation of erectile function would be substantial.

We hypothesize that CN injury-induced apoptosis is caused by decreased SHH protein. A mechanism for SHH involvement in apoptosis induction in the penis is presented in detail below, based on what is known about the Shh signaling pathway in the penis and in other organs (Fig. 9). In the absence of SHH, caspase 3 activity is increased [41, 42] and the G1 to S transition is inhibited, which leads to apoptosis [35, 42] which can be prevented by SHH application. The cell death program is initiated by the proapoptotic dependence receptor PTCH1 [35]. This requires preliminary cleavage of PTCH10s intracellular domain by caspase enzymes (caspase 3 and less frequently caspase 7 and 8 [43]) and exposes the carboxyl-terminal apoptotic domain. Transfecting cultured cells with the carboxy-terminal region of Ptch1 is sufficient to induce cell death [35], increased TUNEL staining is associated with Ptch1 over expression [43], and SHH treatment can inhibit PTCH1-induced cell death in a dose-dependent manner [43]. When SHH is inhibited, PTCH1 inhibits SMO, which causes the formation of GLI3 in a truncated repressor form. Bmp4 is elevated in response to truncated GLI3 during limb development [44] and is associated with elevated apoptosis in several systems [45, 46]. We have shown that SHH protein is decreased following CN injury and that SHH inhibition is able to upregulate BMP4 in the corpora cavernosa [19]. The decrease in SHH protein, accompanied by increased apoptosis, suggests that the mechanism described above may be active in the penis following CN injury. Bmp4 expression was unaltered with CN injury. However, Bmp4 plays a role in and is regulated by several pathways that do not involve Shh, and the protein levels of BMP4 may be different than the RNA expression levels with CN injury, as is the case with SHH. Since SHH treatment is able to prevent CN injury-induced apoptosis in the corpora cavernosa, it is likely that SHH inhibition plays a key role in apoptosis induction in the penis following denervation. However, further experiments are required in order to identify the mechanism of how this process occurs.

FIG. 9.

FIG. 9

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Diagram of the potential role that Shh signaling plays in apoptosis regulation.

A possible mechanism of how SHH maintains penile architecture involves VEGFA. The active form of SHH protein and its receptor PTCH1 are localized in Schwann cells and in veins adjacent to the nerves in the penis. We hypothesize that SHH in Schwann cells of penile nerves regulate homeostasis of adjacent vascular structures through induction of VEGFA. VEGFA acts downstream of SHH in the adult penis, since SHH treatment induces PTCH1 and VEGFA and SHH inhibition downregulates these targets [47]. SHH induction of VEGFA is supported by reports in the literature in cardiac tissue and in ischemic limb [48–50, 42]. Schwann cells have been shown to induce VEGF expression [51, 52]. Mutations that eliminate Schwann cells prevent proper arteriogenesis, and in mutant embryos containing disorganized nerves the trajectory of blood vessel branching is altered to follow the nerve. These data suggest that peripheral nerves provide a template that determines the organotypic pattern of blood vessel branching and arterial differentiation during development, via local secretion of VEGFA. The localization of SHH in Schwann cells of the penis and the ability of SHH to induce VEGFA expression in the penis suggest that a similar mechanism may be active in adult penile nerves and vasculature. Thus SHH may represent a nexus between neural and vascular control of penile morphology.

These studies show that SHH inhibition causes apoptosis in the corpora cavernosa, that morphology changes caused by SHH inhibition in the rat penis are reversible, that CN injury causes a significant decrease in the active form of SHH protein in the corpora cavernosa, and that SHH treatment at the time of CN injury significantly decreases post-CN injury-induced apoptosis. Thus, SHH stabilizes alterations of the corpora cavernosal smooth muscle following CN injury and has significant potential to be developed into a therapy to prevent postprostatectomy apoptosis while the cavernous nerve regenerates.

Footnotes

1

Supported by a grant from the National Institutes of Health/National Institute of Diabetes and Digestive and Kidney Diseases; Grant numbers: DK068507, DK062970, and DK059071.

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