Scaffoldless Tissue Engineering of Stem Cell Derived Cavernous Tissue for Treatment of Erectile Function

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ORIGINAL RESEARCH—BASIC SCIENCE Scaffoldless Tissue Engineering of Stem Cell Derived Cavernous Tissue for Treatment of Erectile Function jsm_2727

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Hazem Orabi, MD,*† Guiting Lin, MD, PhD,* Ludovic Ferretti, MD,*‡ Ching-Shwun Lin, PhD,* and Tom F. Lue, MD* *Knuppe Molecular Urology Laboratory, Department of Urology, School of Medicine, University of California, San Francisco, CA, USA; †Department of Urology, Assiut University, Assiut, Egypt; ‡Experimental Surgical Laboratory, UPRES 4122, Le Kremlin-Bicêtre, France DOI: 10.1111/j.1743-6109.2012.02727.x

ABSTRACT

Introduction. As one-third of erectile dysfunction (ED) patients do not respond to phosphodiesterase-5 inhibitors, there is great demand for new therapeutic options. Adipose tissue-derived stem cells (ADSCs) represent an ideal source for new ED treatment. Aim. To test if ADSCs can be differentiated into smooth muscle cells (SMCs) and endothelial cells (ECs), if these differentiated cells can be used to engineer cavernous tissue, and if this engineered tissue will remain for long time after implantation and integrate into corporal tissue. Method. Rat ADSCs were isolated and differentiated into SMC and ECs. The differentiated cells were labeled with 5-ethynyl-2-deoxyuridine (EdU) and used to construct cavernous tissue. This engineered tissue was implanted in penises of normal rats. The rats were sacrificed after 1 and 2 months; penis and bone marrow were collected to assess cell survival and inclusion in the penile tissues. Main Outcome Measures. The phenotype conversion was checked using morphology, immunocytochemistry (immunohistochemistry [IHC]), and Western blot for SMC and EC markers. The cavernous tissue formation was assessed using rat EC antibody (RECA), calponin, and collagen. The implanted cell survival and incorporation into penis were evaluated with hematoxylin and eosin, Masson’s trichrome, and IHC (RECA, calponin, and EdU). Results. The phenotype conversion was confirmed with positive staining for SMC and EC markers and Western blot. The formed tissue exhibited architecture comparable to penile cavernous tissue with SMC and ECs and extracellular matrix formation. The implanted cells survived in significant numbers in the penis after 1 and 2 months. They showed proof of SMC and EC differentiation and incorporation into penile tissue. Conclusions. The results showed the ability of ADSCs to differentiate into SMC and ECs and form cavernous tissue. The implanted tissue can survive and integrate into the penile tissues. The cavernous tissue made of ADSCs forms new technology for improvement of in vivo stem cell survival and ED treatment. Orabi H, Lin G, Ferretti L, Lin C-S, and Lue TF. Scaffoldless tissue engineering of stem cell derived cavernous tissue for treatment of erectile function. J Sex Med 2012;9:1522–1534. Key Words. Cavernous Tissue; Tissue Engineering; Stem Cells; Adipose-Derived Stem Cells; Erectile Dysfunction Treatment; Cell Sheets

Introduction and the Aim of the Study

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rectile dysfunction (ED) is defined as the persistent inability to attain and maintain penile erection sufficient for sexual intercourse [1]. The Massachusetts Male Aging Study reported a com-

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bined prevalence of minimal, moderate, and complete impotence of no less than 52% [2]. ED leads to significant morbidity and distress, not just for affected men but also for their partners [3]. Therapies for ED include phosphodiesterase type 5 inhibitors (PDE5Is), intracavernosal vasoactive © 2012 International Society for Sexual Medicine

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Tissue Engineering of Cavernous Tissue injections, and vacuum erection devices, penile implants [4]. Despite the overwhelming success of PDE5I, the demand for pharmacotherapeutic and surgical options for ED continues to rise as a result of the increased proportion of elderly in the population in addition to the growing percentage of ED patients who do not respond to PDE5I [5]. Surgical treatment of ED is associated with many complications, which warrants the need for nonsurgical therapies for those patients [6]. In addition, none of the above treatments truly correct, cure, or prevent ED. These inadequacies have led many scientists and clinicians to investigate new therapies, mainly gene therapy and regenerative medicine. Gene therapy, although promising, has many challenges and obstacles like local inflammatory response and random transgene expression, beside other safety issues that limit its use at clinical level [7,8]. With regard to the regenerative medicine, there are two main approaches. The first one includes the regeneration of the corpora cavernosa with the use of autologous cells seeded on scaffolds [9–11]. However, many challenges are in face of this biotechnology as fulfillment of architectural, biochemical, and functional requirements of native corpora cavernosa and complexity of connecting these grafts to functional blood circulation and autonomic innervation. The second one involves cell-based therapy with the injection of functional or stem cells (SCs) into the corpus cavernosum. This approach is more practical as it avoids reconstruction of all penile elements and architecture. Cellular therapy includes the use of endothelial progenitor cells (EPCs), muscle-derived SCs, and embryonic and mesenchymal SCs [12–17]. EPCs show some phenotypic overlaps with hematopoietic cells and debate still exists with respect to the identification and origin of EPCs [18]. In addition, patients with ED have a lower number of circulating EPCs probably as a result of a generalized endothelial dysfunction [19]. Muscle-derived SCs have the disadvantages of biopsy problems and differentiation into only smooth muscle cells (SMCs), not endothelial cells (ECs), whose dysfunction is a major pathology in ED. The practical use of embryonic SCs is limited due to ethical considerations, malignant potential, and problems with cell regulation. Although bone marrow (BM)derived SCs have many advantages, they encompass the problems of inaccessibility, traumatic collection, and small heterogeneous yield of SCs. Adipose-derived SCs (ADSCs) represent a potential solution for cellular therapy of ED. We

reported the improvement of the erection in experimental diabetic, hyperlipidemic, and cavernous nerve injury models after intracavernous (IC) injection of unmodified ADSCs [16,17,20]. However, there was paucity of ADSCs remaining in the corpus cavernosum and the number of differentiated cells after 4 weeks. We are exploring new approach to improve the durability of cell residence and integration of injected cells into the penile tissue. This approach depends on scaffoldless formation of cavernous tissue from ADSCs in vitro and implanting it into the penis. This study is designed to test if the ADSCs can be differentiated into SMC and ECs, if these differentiated cells can be used to engineer scaffoldless cavernous tissue in vitro and deliver it to the rat penis, and if this engineered cavernous tissue cells will remain for long time after its implantation and integrate into corporal tissue. Materials and Methods

A study overview is shown in Figure 1A. The total number of animals used for the entire study was 18:9 animals, which were euthanized 1 month after in vivo SC implantation, and nine animals were euthanized 2 months after in vivo SC implantation. The age of the animals was 3 months at the time of harvest of ADSCs.

Harvesting ADSCs All animal experiments complied with National Institute of Health guidelines and were approved by the Institutional Animal Care and Use Committee at the University of California, San Francisco. All animals were Sprague-Dawley rats purchased from Charles River Laboratories (Wilmington, MA, USA). After adequate induction of anesthesia, a midline laparotomy was performed and perigonadal fat pad was identified. The fat pad was sharply excised unilaterally with hemostasis. The fat pad was placed in phosphate buffered saline (PBS) for cell culture later. The abdomen was closed in two layers. Cell Isolation, Expansion, and Differentiation Cell harvesting, growth, and expansion were carried out in a well-equipped cell culture facility in our laboratory. ADSCs were harvested according to known protocol established in our laboratory using collagenase type IA. The tissue was incubated with collagenase IA (Sigma-Aldrich Corp., St. Louis, MO, USA) for 1 hour at 37°C with vigorous shake for 15 seconds in 20-minute intervals. After J Sex Med 2012;9:1522–1534

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Figure 1 Showing study design and surgery of penile implantation. ADSC = adipose-derived stem cell; EC = endothelial cell; EdU = 5-ethynyl-2-deoxyuridine; SMC = smooth muscle cell; SMER = smooth muscle endothelial reconstruct

centrifugation, the resulting pellet, which is defined as the adipose tissue stromal-vascular fraction (SVF), was exposed to lysis buffer for 10 minutes to remove red blood cells. The remaining cells were filtered through 40-mm cell strainer, suspended in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 1% nonessential amino acid, 1% penicillin, streptomycin fungizone (PSF) (10,000 units/mL penicillin, 10,000 mcg/mL streptomycin SO4, and 0.025 mg/mL fungizone), and 110 mg/mL sodium pyruvate, and plated at a density of 1 ¥ 106 cells per dish. The next day, the culture medium was disJ Sex Med 2012;9:1522–1534

carded by aspiration and the cells washed with PBS, and cells were differentiated toward either SM lineage using DMEM supplemented with 10% FBS with transforming growth factor-b1 (TGFb1) (2 ng/mL TGF-b1 [R & D Systems Inc., Minneapolis, MN, USA]) or EC type using endothelium growth medium-2 (EGM-2) medium supplemented with growth factors and cytokines (Lonza Biologics Inc., Allendale, NJ, USA). The cell differentiation toward SMC was checked with immunohistochemistry (IHC) using SM actin (SMA), calponin and SM myosin heavy chain antibodies, and Western blot. The EC differentiation

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Tissue Engineering of Cavernous Tissue was confirmed with IHC using rat EC antibody (RECA) and von Willebrand factor (vWF) antibody, low-density lipoprotein (LDL) uptake, and Matrigel-based capillary-like tube formation assay. After that, the cells were labeled using 5-ethynyl-2-deoxyuridine (EdU) (Invitrogen, Carlsbad, CA, USA) to track the cells in vivo after penile implantation.

In Vitro Cavernous Tissue Formation Induced SMCs were plated for 4–6 days on specially coated dishes, and then ECs were seeded on top for additional 2 days to form cavernous tissue. These dishes are thermosensitive 3.5 cm with Nunc UpCell Surface, purchased from ThermoScientific Nunc (Cat. no. 174904; Rochester, NY, USA). The cavernous sheet was detached by lowering the temperature below 20°C. Part of the cavernous tissue was fixed and frozen in optimum cutting temperature compound (Sakura Finetek, Torrance, CA, USA) and stored at -80°C until use. Sections were cut at 5 mm, adhered to charged slides, air dried for 10 minutes, and rehydrated with PBS. The section was stained for Masson’s trichrome staining IHC with antibodies against RECA, calponin, and collagen IV. Intracorporal Delivery of the Cavernous Tissue The penile skin was incised to reveal the rat corporal bodies. A small incision was done in the tunica albuginea of the proximal part of the penis on one side, and the cavernous tissue was placed under tunica. A single 6-0 nylon suture was placed to close the tunical incision and mark the implantation site. This was followed by closure of the penile skin with 3/0 vicryl suture (Figure 1B). The animals were scarified at 1 and 2 months after implantation, and the penis and BM were collected for histological evaluation. The histological assessment included biochemical staining (hematoxylin and eosin [H&E] and Masson’s trichrome) and IHC (RECA, calponin, and EdU). Image Analysis The stained tissues were examined with a Nikon Eclipse E600 fluorescence microscope (Nikon Instruments, Melville, NY, USA) and photographed with a Retiga 1300 Q-Imaging camera (Q-Imaging, Surrey, BC, Canada) using the ACT-1 software (Nikon Instruments). Computerized histomorphometric analysis was performed using Image-Plus 5.1 software (Media Cybernetics, Bethesda, MD, USA).

Procedures

Immunofluorescence for Cells Cells were seeded onto a coverslip inside each well of a six-well plate at 40–60% confluence in DMEM with 10% FBS in case of induced SMC and EGM-2 in case of induced ECs for 24 hours. The next day, the cells were fixed with ice-cold methanol for 5 minutes, permeabilized with 0.05% triton X-100 for 8 minutes, and then blocked with 5% normal horse serum in PBS for 1 hour at room temperature. The cells were then incubated with mouse SMA (Sigma-Aldrich Corp.; 1:2,000), anticalponin (Santa Cruz Biotechnologies, Santa Cruz, CA, USA; 1:500), anti-SM myosin (Abcam Inc., Cambridge, MA, USA; 1:500), anti-vWF antibody (Abcam Inc.; 1:500), and mouse anti-RECA (Santa Cruz Biotechnologies; 1:500) for 1 hour. Then, the cells were incubated with 1:500 dilution of secondary antibody conjugated with Alexa fluor 488 (Invitrogen) or Alexa fluor 594 (Invitrogen) for 1 hour. After three rinses with PBS, the cells were stained with 4′,6diamidino-2-phenylindole (DAPI, Sigma-Aldrich Corp.) for 5 minutes. LDL Uptake Cells were seeded into six-well plates in DMEM (in case of ADSC as control) or EGM-2 medium (in case of induced EC) and incubated at 37°C. The next day, 10 mg/mL of acetylated LDL DiI complex (DiI AcLDL, Invitrogen) was added to the culture medium. The next day, the cells were examined by phase-contrast and fluorescence microscopy, and photographed. Matrigel-Based Capillary-Like Tube Formation Assay Six-well tissue-culture plate was plated with 300 mL of growth factor-reduced Matrigel (BD Biosciences, San Jose, CA, USA) per well. Approximately 5 ¥ 104 cells in 500 mL of EGM-2 were then seeded into each well and incubated at 37°C. Twenty-four hours later, development of capillary-like networks was examined by phasecontrast microscopy and photographed. Human umbilical vein ECs served as a positive control. Immunofluorescence for Animal Tissues After euthanasia, tissue samples were fixed in cold 2% formaldehyde and 0.002% picric acid in 0.1 M phosphate buffer, pH 8.0, for 4 hours followed by overnight immersion in buffer containing 30% sucrose. Then, the tissues were frozen in optimal cutting temperature tissue (OTC) and stored at J Sex Med 2012;9:1522–1534

1526 -80°C until use. Sections were cut at 5 mm, adhered to charged slides, air dried for 10 minutes, and rehydrated with PBS. Goat serum 3% in PBS was applied as blocking agent for 30 minutes. Sections were incubated overnight at 4°C with primary antibodies, followed by 1-hour incubation in 1:500 dilution of secondary antibody conjugated with Alexa fluor 488 (Invitrogen) or Alexa fluor 594 (Invitrogen). Primary antibodies were anticalponin (Santa Cruz Biotechnologies; 1:500), mouse anti-a-SMA (Sigma-Aldrich Corp.; 1:2,000), mouse anti-RECA (Santa Cruz Biotechnologies; 1:500), and anticollagen (Abcam Inc.; 1:500). Nuclear staining was performed by 2-minute incubation in DAPI (D-3571, Invitrogen). For tracking of ADSC with EdU, slides were incubated with freshly made Click-iT reaction cocktail, which contained Alexa-594 fluor (Invitrogen) for 30 minutes at room temperature. Nuclear staining was performed with DAPI (D-3571, Invitrogen). Results

Endothelial Differentiation in EGM-2 ADSCs were isolated in efficient way each time. The yield of cell in each time is considerable in amount and sufficient for each animal. We routinely culture ECs in EGM-2, which is a commercially available endothelial growth medium. When DMEM in ADSC cultures was replaced with EGM-2, the cells reached confluence faster and appeared more compact (larger nuclei) than cells that remained in DMEM with characteristic cobble stone appearance of ECs. IHC showed that induced ECs expressed endothelial-specific markers RECA and vWF. LDL uptake assay showed that ADSCs grown in EGM-2 were capable of LDL uptake. Additionally, Matrigel tube formation assay beside LDL test also showed that ADSCs grown in EGM-2 were able to form endothelial-like tube structures. The endothelial specificity of these three assays was supported by positive results with human umbilical vein endothelial cells and negative results with ADSC grown in DMEM (Figure 2). SM Differentiation Prior to culture with the differentiating media, ADSC did not express calponin or SM myosin, however, low expression of SMA was seen with IHC. After induction of SMC differentiation, J Sex Med 2012;9:1522–1534

Orabi et al. there was progressive increase in the expression of these SMC markers overtime. Western blot analysis after 3 weeks (Figure 3) revealed up-regulation of SMA protein. Spindle shape of the induced ADCS was fully achieved after 3 weeks.

Cavernous Tissue Formation After 6–8 days, multilayer tissue was formed and could be detached by lowering the temperature below 20°C. The upper cell layer should be positive staining for RECA, followed by a layer of collagen IV and then few cell layers showing positive staining for calponin. This indicated that the cavernous tissue is formed of upper layer of ECs, extracellular matrix (ECM), and SMCs. This cavernous tissue was called SM endothelial reconstruct (SMER). SMER is thin, yellowish in color, and can be handled with fine surgical instruments (Figure 4). In Vivo Implantation All animals tolerated the surgery without complications. Autologous EdU-labeled ADSCs were transplanted into corpus cavernosum and identified by H&E, Masson’s trichrome, and Alexa-594 and 488 stains (Invitrogen, Molecular Probes, Invitrogen, Grand Island, NY, USA). The histological data indicated the presence of EdU-labeled cells in the corpus cavernosum in significant amounts after 1 and 2 months in comparison to our previous studies. They were distributed near the implantation site after 1 month; however, they were distributed in corpus cavernosum after 2 months (Figure 5). Mild increase in the collagen deposition was observed at the site of implantation. This may be due to trauma of tunica albuginea or new collagen formation by the implanted cells. The EdU-labeled cells exhibited endothelial or SMC differentiation or no differentiation. The induced ADSCs formed capillaries, contributed to cavernous sinusoids, or stay differentiated but nonintegrated (Figure 6). The induced SMC differentiation was less evident and induced cells were either incorporated into penile vasculature or stay nonintegrated in the cavernous tissue (Figure 7). Discussion

Ideally, a SC for replacement therapy should be found in abundant quantities, harvested by a minimally invasive procedure, and differentiated along multiple cell lineage pathways in a regulatable and

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Figure 2 Showing endothelial differentiation (ECs) of ADSCs as confirmed with IHC using RECA (A), LDL uptake (B), and tube formation on Matrigel (C). ADSC = adipose-derived stem cell; EC = endothelial cell; HUVEC = human umbilical vein endothelial cell; IHC = immunohistochemistry; LDL = low-density lipoprotein; RECA = rat endothelial cell antibody

reproducible manner. Also, it can be safely and effectively transplanted to either an autologous or allogeneic host and manufactured in accordance with current Good Manufacturing Practice guidelines [21]. For these reason, ADSCs represent an ideal source for SC therapy [22]. Additionally, ADSCs are likely vascular SCs at various stages of differentiation toward becoming SM and ECs [23]. Furthermore, ADSCs have the ability to secret many multiple potentially synergistic proangiogenic growth factors delineating their

angiogenic and antiapoptotic potential which are key therapeutic factors in treatment of ED [24]. In different types of ED, ECs, cavernous SMCs (CSMCs), and cavernous nerves are often altered and rendered apoptotic. As a result, transplantation of SC into penis should replenish the depleted EC and/or CSMC pools. However, in published preclinical studies, the cellular differentiation occurred in the treated animals is controversial and SC contribution into penis on long term was poor [16,17,20]. Consequently, paracrine actions J Sex Med 2012;9:1522–1534

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Figure 3 Showing smooth muscle cell differentiation of ADSCs as confirmed with IHC using SMA, calponin and SM myosin, and Western blotting with SMA. ADSC = adipose-derived stem cell; IHC = immunohistochemistry; SM = smooth muscle; SMA = smooth muscle actin

as opposed to cellular differentiation were responsible for SC’s therapeutic efficacy in these studies. This may limit treatment duration and efficacy. The importance of cellular differentiation and residence to durable and efficient therapeutic effect was also suggested in a clinical trial using umbilical cord blood SCs [25]. Despite having increased penile rigidity, none of the patients was able to achieve vaginal penetration unless aided by taking sildenafil before coitus. During 11-month follow-up, only one treated subject maintained erection sufficient for coitus. The authors supposed that the possibility of transdifferentiation of grafted SCs might be responsible for the maintenance of satisfactory erection until at least 11 months after treatment. They also suggested that changing patterns of effects from early increase followed by later decline can be explained by cell numbers beside other factors. J Sex Med 2012;9:1522–1534

To overcome these hurdles, we explored a novel way to improve the cellular differentiation, cell residence, and engraftment. ADSCs were differentiated into ECs when grown in EGM-2, which did contain factors necessary and sufficient to induce ADSC endothelial differentiation. We previously demonstrated that ability [26]. Boosting the corpus cavernous with ECs will help to overcome the endothelial dysfunction which is a key pathology in ED. ADSCs were also differentiated into SMC when grown in differentiation media. We confirmed that induced ADSCs showed variable expression of SMC molecular markers confirming their phenotype conversion. The cells acquired spindleshaped appearance after 3 weeks in differentiation media. The expression of SMC markers was successive starting with SMA, then calponin, and lastly SM mysosin as evidenced by IHC and

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Figure 4 Showing smooth muscle endothelial reconstruct (SMER) in vitro composed of ECs, SMC, and collagen as evidenced with IHC (A,B) with RECA, collagen IV, and calponin antibodies. SMER is seen detached from the culture dish in (C) section. DAPI = 4′,6-diamidino-2-phenylindole; EC = endothelial cell; IHC = immunohistochemistry; RECA = rat endothelial cell antibody; SMC = smooth muscle cell

Western blot. Our result was consistent with Harris et al. who proved that the use of high concentration of FBS with other molecules including angiotensin II, sphingosylphosphorylcholine, or TGF-b1 led to SMC differentiation of ADSCs [27]. More interestingly, Basu et al. isolated a cellular subpopulation of adipose SVF that is separate and distinct from other classes of adipose-derived cells after expanding SVF cells with DMEM plus 10% FBS. They were called adipose-derived SM-like cells as they consistently express SMC markers, independent of donor site and across multiple passages [28]. This confirms the tendency of ADSCs to be easily differentiated with high percentage into SM and ECs. Differentiated cells have a superior graft survival, early and better cell integration, and engraftment into host tissues. Additionally, cellular differentiation is desirable as it is suggested to prevent ectopic tissue and tumor formation especially in the light of discovering aberrant cell line of ADSCs with tumor potential [29]. The principal route for SC transplant is intracorporal injection which is systemic in nature. In all of our published SC-for-ED studies, we reported difficulties in finding the transplanted SC in penile [12,16,17,20]. In studies published by others, IC injected SC was similarly difficult to find. In our recent studies, majority of IC injected SC exited

the penis within 1 day [30,31]. To overcome this problem, we imposed the idea of anatomically focused cellular therapy by creating new technology called SM endothelium reconstruct (SMER) simulating normal cavernous sinusoidal tissue. The formed tissue was detached by lowering the temperature resulting in continuous cell sheet in a reproducible and consistent all over the study. Being thin and easily handled surgically, SMER has the advantage of many routes of delivery including surgical implantation and IC injection. It is composed of upper layer of ECs and multiple layers of SMCs with ECM composed mainly of collagen in between. Collagen IV is found primarily in the basal lamina [32]. The presence of the ECM allowed the attachment and final maturation of both ECs and SMCs. Also, the presence of the cells in this form prevented their wash out by the circulation in case of IC injection and helped fixation of the tissues to the site of implantation. Also, own ECM and detachment by lowering the temperature avoided the trypsinization process. Trypsinization interrupts interactions between cultured cells and their extracellular matrices, facilitating apoptosis and consequently limiting the therapeutic efficacy of the transplanted cells. The implantation process was easily done without any recorded complications. The data showed that implanted cells were identified in J Sex Med 2012;9:1522–1534

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Figure 5 Showing the local retention of the implanted cells near the site of the implantation after 1 month as shown by H&E and Masson’s trichrome and EdU staining (A). After 2 months, there is still retention and survival of significant number of the implanted cells (B). EdU-labeled ADSCs migrated to BM after 1 month (C). ADSC = adipose-derived stem cell; BM = bone marrow; DAPI = 4′,6-diamidino-2-phenylindole; EdU = 5-ethynyl-2-deoxyuridine; H&E = hematoxylin and eosin; SMER = smooth muscle endothelial reconstruct

penis 1 and 2 months after implantation as verified by H&E, Masson’s trichrome staining, and EdU labeling. One month after postoperatively, dense collection was seen at implantation site. However, by 2 months, many cells were seen dispersing throughout the corpus cavernosum although smaller ADSC cell collection remained nearest the implantation site. The cells were implanted in the proximal part of the corpus cavernosum and were detected in both the proximal and distal parts in the same corpus after 2 months. However, we did not detect the ADSCs in the unimplanted corpus. This may be due the short time interval (only 2 months) or lack of actual distribution of cells into the unimplanted corpus. The difference between J Sex Med 2012;9:1522–1534

H&E and Masson’s trichrome staining and EdU staining may be attributed to the long duration of the in vitro cell culture prior to their transplantation (6–8 days) and the long time of implantation as its EdU label gets diluted with each round of cell division. The cell distribution can be explained as SMER was fixed to site of implantation early postoperative and later parts of SMER were detached and cells spread within the corpus cavernosum. Improved local retention and net survival of the transplanted cells could lead to improvement in the therapeutic effect of the cells and, hence, greater improvement in erectile function after treatment. This also would lead to prolongation and magnification of paprcrine action of the

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Figure 6 Showing the expression of the implanted ADSCs for the phenotype of endothelial cells as evident by positive staining for RECA near and away from the site of implantation. Many endothelial cells remained nonintegrated in the corporal tissue (A). EdU-labeled endothelial cells were incorporated in cavernous sinusoids (B) while others were seen forming blood capillaries (C). ADSC = adipose-derived stem cell; DAPI = 4′,6-diamidino-2-phenylindole; EdU = 5-ethynyl-2-deoxyuridine; RECA = rat endothelial cell antibody

Figure 7 Showing the expression of the implanted ADSCs for the phenotype SMC phenotype as evident by positive staining for calponin. The cells remained nonintegrated (A) or incorporated in penile vasculature (B). ADSC = adipose-derived stem cell; DAPI = 4′,6-diamidino-2-phenylindole; EdU = 5-ethynyl-2-deoxyuridine; SMC = smooth muscle cell

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1532 transplanted cells. The use of bioabsorable biomaterials and scaffolds is a well-known method of SC delivery to improve local cell retention graft survival and possibly therapeutic effect [33–35]. However, this technique has the disadvantages of antegenicity, complexity of procedure, incomplete cell-biomaterial attachment and interaction, and possible hazards and complications. Our form of SC delivery (SMER) avoids the use of scaffolds or biomaterials as method of SC delivery because the induced ADSCs make their own ECM that helps better cell residence after transplantation. In addition, our new technology provides the cells in an ideal form where all layers of the cavernous tissue are present which offers better cell survival and engraftment. Some of labeled transplanted cells were found in BM as a number of cells detached and migrated through blood stream to BM. This is supported by the fact that corpus cavernosum is composed of endothelium-lined sinusoids that are anatomically and physiologically similar to arteries and veins [36]. Thus, IC injection is analogous to intravenous injection in that the injected cells can be transported by blood to distant sites, including the target tissue. This has been proved in one of the latest publications where ADSCs exited the penis and traveled preferentially to BM within days of IC injection [30]. This trait in addition to cellular differentiation prior to their transplantation adds to the safety of the use of ADSCs in treating ED. EC dysfunction has been proposed as an important mechanism in ED especially diabetic-induced ED [37,38]. Replacing the lost and dysfunctional ECs would be of paramount importance in restoring the erectile function. In previous studies, although the IC injection of unmodified ADSCs improved ED, however, the number of differentiating ADSCs into ECs was scanty [12,16,17,20]. In our study, there was increased number of ADSC-derived ECs in the corpus cavernosum. This can be attributed to the implantation of differentiated ECs and the longterm withholding of implanted ADSCs. Boosting the penis with new ECs is expected to improve the therapeutic effect of the ADSCs. These ECs formed blood capillaries, contributed to cavernous sinusoids, and remained nonintegrated. The explanation why many of the ECs remained nonintegrated would be the fixation of cells of SMER by adhesions due to its site of implantation early postoperative. This can be avoided later by embedding SMER deeply into the cavernous tissue, not under the tunica. J Sex Med 2012;9:1522–1534

Orabi et al. EdU-labeled SMCs were found in the corpus cavernosum. This was in accordance with our previous studies when ADSCs were injected into the penis and with other studies involving the injection of human ADSCs into the bladder and urethral sphincter [39]. EdU-labeled SMCs were not detected in significant amount due to the fact that the induced SMCs need to be implanted between SMs to keep their phenotype and function, which was not the case in our study as the SMER was implanted below the tunica where fibroblasts predominate. The microenvironment provides growth factors and other signals that guide appropriate differentiation of transplanted cell populations [40]. Part of the implanted cells did not show positive staining to either SMCs or endothelial markers. These cells either lost differentiation capacity and protein expression and returned to their steaminess or transformed into other cell types, mostly fibroblasts, because of implantation site. This will be investigated in our next studies. This is important as these SCs will act as depot for reparative SCs, which is affected in aged and diabetic ED. We did not compare implanting vs. injecting SCs into the penis at the same study environment as the following were the same: method of adipose SC harvest, age of rats, growth medium, cell culture dishes, laboratory environment, and technique as reported in the previous studies (with the exception of those used in differentiation and SMER formation). More importantly, in one of our recent publications, when ADSCs (unmodified) were injected into the corpus cavernosum of healthy rats, we demonstrated that ADSCs exited the penis and could not be found in penile tissues 4 weeks later [31]. Limitations to our study include the lack of diseased animal model and functional improvement after the implantation of SMER. Correspondence Author: Hazem Orabi, MD, Assiut University, Urology, Urology Department, Asiut University Hospitals, Assiut 71516, Egypt. Tel: +20100454008; Fax: +2088233327; E-mail: hazem. [email protected] Conflict of Interest: None.

Statement of Authorship

Category 1 (a) Conception and Design Hazem Orabi; Tom F. Lue (b) Acquisition of Data Hazem Orabi; Guiting Lin; Ludovic Ferretti

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Tissue Engineering of Cavernous Tissue (c) Analysis and Interpretation of Data Hazem Orabi; Guiting Lin; Ching-Shwun Lin; Tom F. Lue

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Category 2 (a) Drafting the Article Hazem Orabi; Tom F. Lue (b) Revising It for Intellectual Content Hazem Orabi; Tom F. Lue

Category 3

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