Smooth Muscle Insulin Receptor Deletion Causes Voiding Dysfunction: A Mechanism for Diabetic Bladder Dysfunction

Insulin signaling is crucial for glucose metabolism and protein/lipid synthesis, and recent progress indicates that it is also important for gene transcription, cell proliferation, and cell differentiation, in regulating multiple pathways, including mTOR and FoxO signaling (1–8). Studies in several organs suggest that insulin signaling plays differential roles in different tissues by regulating a variety of downstream effectors. For example, specific IR deletion in skeletal muscle results in increased serum triglycerides and decreased muscle growth (4,9), and liver-specific deletion creates severe insulin resistance associated with glucose intolerance (4,10), while in adipose tissue, deletion of IR markedly reduced fat mass and protected the animal from age-related obesity and glucose intolerance (11).

Diabetic bladder dysfunction (DBD) is the most common complication of diabetes, affecting 40–100% of patients with diabetes (12). DBD is commonly associated with myogenic and neurogenic alterations. Patients often exhibit decreased sensation, polyuria with increased bladder volume, poor emptying with increased postvoid residual urine volume, and bladder instability and overactivity, as well as urinary urgency and incontinence (13–20). These debilitating symptoms of DBD significantly impact quality of life.

The control of bladder smooth muscle (BSM) contractility is central to whether the bladder functions properly. In general, micturition is mediated by neurotransmitters ATP and acetylcholine (Ach), which are coreleased by parasympathetic nerves. ATP binds to P2x1 receptor and elicits a rapid contraction to initiate voiding, while Ach binds to muscarinic M₂/M₃ receptors to induce a slower cholinergic contraction to expel urine (21–25). Indeed, alterations in BSM contractility have consistently been observed in DBD of both people and animal models (13–20). With induction of diabetes with streptozotocin in animal DBD studies and a Zucker diabetic fatty (ZDF) rat model, diabetes caused significantly altered BSM contractility in response to electric field stimulation (EFS), KCl, or agonists like carbachol and α,β-methyleneadenosine 5′-triphosphate trisodium (α,β-meATP) (20,26).

The molecular mechanisms underlying DBD are not fully understood. Hyperglycemia-induced polyuria might play a role in bladder hypertrophy and overactivity, and oxidative stress due to hyperglycemia might induce molecular and cellular damage of bladder tissue (27). As mentioned, insulin signaling might play a central role in the pathogenesis of DBD, and injection of insulin has been shown to reverse streptozotocin-induced DBD in animal models (17). A recent study has shown that deletion of insulin receptor in arterial smooth muscle significantly reduces intimal hyperplasia in a high-fat-fed mouse model, suggesting insulin signaling is an important pathway during the pathogenesis of restenosis in diabetes (28). How insulin receptor signaling in bladder tissue regulates bladder function and whether the disruption of insulin signaling in bladder tissue itself contributes to the pathogenesis of DBD are, however, completely unknown. To address this question, we generated both heterozygous and homozygous smooth muscle–specific IR knockout mouse models to mimic the disrupted insulin signaling in diabetes. These mice exhibited altered voiding function, abnormal bladder morphology, impaired BSM contractility, and dysregulated IR-mediated signaling in bladder, which suggests a pathogenesis recapitulating DBD in human patients with diabetes.

Unless otherwise specified, all chemicals were obtained from MilliporeSigma and were of reagent grade or better. P2x1 agonists α,β-meATP and carbachol were purchased from R&D Systems.

All mice used in this study were on the C57BL/6J background and 12–16 weeks old with matching conditions for age and sex. Both male and female mice were used unless otherwise specified. The mice were housed in standard polycarbonate cages with free access to normal food and water. Wild-type C57BL/6J mice were purchased from The Jackson laboratory (Bar Harbor, ME). We generated both heterozygous and homozygous smooth muscle–specific insulin receptor–deficient mice (SMIR^+/− and SMIR^−/−) by mating INSR^f/f mice (generously donated by Dr. C. Ronald Kahn’s Laboratory at Harvard Medical School) with Sm22α-cre mice (B6.Cg-Tg(Tagln-cre)1Her/J; The Jackson Laboratory). All animal studies were performed in adherence with U.S. National Institutes of Health guidelines for animal care and use and with the approval of the Beth Israel Deaconess Medical Center Institutional Animal Care and Use Committee.

Glucose tolerance tests (GTTs) were performed on mice fasted overnight (14 h), and then glucose (2 mg/g body wt i.p.) was administered. Insulin tolerance rests (ITTs) were performed on mice fasted for 6 h, and then insulin (0.75 mIU/g body wt i.p.) was administered. Blood glucose was measured from tail nicks at 0, 15, 30, 45, 60, and 120 min after glucose or insulin injection by Contour glucometer (Bayer, Mishawaka, IN).

Mice fasted for 14 h were injected with glucose (2 mg/g body wt i.p.). Blood was sampled from the tail at 0, 15, 30, 45, 60, and 120 min after glucose injection. Plasma insulin levels were measured with a mouse insulin ELISA kit (Ultrasensitive Mouse Insulin ELISA kit, cat. no. 90080; Crystal Chem) on an xMark Microplate Spectrophotometer (Bio-Rad Laboratories) according to the manufacturer’s instructions.

Void spot assay (VSA) was performed during daytime with light from ∼9:00 a.m. to 1:00 p.m. in female mice. We did not perform VSA on male mice because male voiding data are often complicated by their territorial marking behavior. Individual mice were gently placed in a standard mouse cage with BLICK Cosmos Blotting Paper (cat. no. 10422-1005) placed at the bottom, during which time water was withheld and standard dry mouse chow was available. Recovered filters were imaged as previously described (29,30). Overlapping voiding spots were visually examined and manually separated by outlining and copying and then pasting to a nearby empty space in Fiji software for analysis. A volume:area standard curve showed that a 1 mm² voiding spot represents 0.283 μL urine. Urine spots with an area ≥80 mm² were considered primary voiding spots (PVS) based on a cutoff established from the voiding spot patterns analyzed from hundreds of mice (29–31).

BSM myography was performed on male mice. Male have larger bladders (∼30 mg) compared with female mice, which is easier for dissecting away the epithelial layer and yields four muscle strips per bladder consistently. Mucosa-dissected BSM strips were cut longitudinally (∼2 mm wide and ∼7 mm long). Muscle strips were mounted in an SI-MB4 tissue bath system (World Precision Instruments, Sarasota, FL). Force sensors were connected to TBM4M Transbridge (WPI), and the signal was amplified by PowerLab (ADInstruments) and monitored with LabChart software (ADInstruments). BSM strips were gently prestretched to optimize contraction force and then pre-equilibrated for at least 1 h. Contraction force was sampled at 2,000/s with use of LabChart software. EFS was carried out with a Grass S48 field stimulator (Grass Technologies, West Warwick, RI) according to previously described standard protocols: voltage 50 V, duration 0.05 ms, trains of stimuli 3 s, and frequencies 1, 2, 5, 10, 20, and 50 Hz (32,33).

Excised bladders were fixed in 4% formaldehyde, embedded in paraffin blocks, and the middle equatorial region of the bladder was sectioned prior to staining with Masson trichrome for bladder wall thickness evaluation. A minimum of six points (shortest distance) where infolding urothelial layers were lacking were averaged on each section and three or more sections per bladder, and the final mean is bladder wall thickness. Muscle layer thickness is measured on six or more randomly chosen points on each section, and three or more sections per bladder. For immunofluorescence staining, excised bladders were fixed in 4% (w/v) paraformaldehyde and then cryoprotected, frozen, sectioned, and incubated with antibodies (1:100) (Supplementary Table 1) overnight at 4°C. The sections were then incubated with an Alexa Fluor 488–conjugated secondary antibody (diluted 1:100), and nuclei were stained with DAPI. Imaging was performed on an Olympus BX60 fluorescence microscope with a 40×/0.75 objective or on a Leica THUNDER imaging system with a 63× objective (33).

Whole bladder tissues from female mice were lysed with radioimmunoprecipitation assay buffer (50 mmol/L Tris-HCl, 150 mmol/L NaCl, 1% NP-40, 0.5% sodium deoxycholate, and 0.1% SDS). The protein concentration was determined with use of BCA protein assay reagent (Thermo Fisher Scientific, Rockford, IL). Proteins (25 µg) were loaded into each well and separated by SDS-PAGE and transferred to polyvinylidene fluoride membranes. Membranes were incubated with primary antibodies (Supplementary Table 1) at 4°C for 2 h. Protein bands were visualized with Amersham ECL reagent (Arlington Heights, IL). The membranes were incubated with Restore PLUS Western Blot Stripping Buffer (Thermo Fisher Scientific) for 5 min to remove primary and secondary antibodies for reblotting with new antibodies, and each membrane was stripped and reprobed no more than three times. The intensity of protein bands was scanned and then quantitated with Fiji software (33). Targeted protein in each lane was normalized to β-actin intensity in the same lane. The average protein expression level in wild-type mice was designed as 1, and the fold change of protein expression level in knockout mice was reported.

All data are presented as box and whisker plots. The centerline is the median, the box represents 75% of the data, and the whiskers indicate minimum to maximum. Data were analyzed with one-way ANOVA for comparison among groups. Bonferroni multiple comparisons post hoc tests were used where necessary between the two groups with GraphPad Prism 8 software, and a P value <0.05 was considered significant.

Further information, reagents, and other supporting data in this study are available from the corresponding author on request.

Given that IR plays a crucial role in glucose homeostasis, glucose metabolism in SMIR^+/− and SMIR^−/− mice was assessed. In comparisons with wild-type mice, there was no detectable difference in blood glucose levels in both nonfasted and fasted female SMIR^+/− and SMIR^−/− mice (Fig. 1A). Male mice were similar (Supplementary Fig. 1A). However, both male and female SMIR^−/− mice exhibited hyperglycemia when challenged with an injection of glucose (2 mg/g body wt i.p.). There was elevated blood glucose in SMIR^+/− mice in response to GTT, but the difference was not statistically different compared with wild type (Fig. 1B and C and Supplementary Fig. 1B and C). Consistent with the GTT response, blood glucose level in both male and female SMIR^−/− mice in response to ITT was significantly higher than in wild-type animals (Fig. 1D and E and Supplementary Fig. 1D and E), indicating impaired ability of SMIR^−/− mice in glucose metabolism during stressed condition. In comparison with wild-type mice, the plasma insulin levels were not different in either SMIR^+/− or SMIR^−/− female mice. The overall plasma insulin levels in response to GTT were also similar in all three studied groups, though the SMIR^−/− mice tend to show mildly elevated insulin levels (Fig. 1F–H).

For understanding of whether deficiency or deletion in BSM insulin signaling can directly cause voiding dysfunction, we performed VSA to examine the voiding phenotype in both SMIR^+/− and SMIR^−/− mice. Compared with wild-type mice, which produce on average 3.03 PVS in a 4-h period, significantly increased voiding frequency was seen in SMIR^+/− (5.83 PVS/4 h) and SMIR^−/− (5.07 PVS/4 h) mice. Wild-type mice have an average PVS size of ∼434 mm²/spot (∼123 μL/void); however, PVS size is significantly smaller in SMIR^+/− (319 mm²/spot, ∼90.33 μL/void) and SMIR^−/− (273 mm²/spot, ∼77.29 μL/void) mice (Fig. 2A–F). Voiding spots were further grouped according to their different sizes as shown in the frequency distribution chart in Fig. 2G, and spots showed a distribution pattern similar to a skewed bell curve, with control group peaks at ∼200–300 mm², while SMIR^+/− and SMIR^−/− groups peak at ∼80–200 mm², indicating a significantly shifted voiding pattern. For confirmation of the altered voiding pattern detected in VSA and exploration of potential underlying mechanisms, urodynamic studies were performed with CMG. CMG data consistently indicated that SMIR^+/− and SMIR^−/− mice exhibited increased voiding frequency with decreased bladder compliance (Fig. 2H–N).

To understand how deficiency in BSM insulin signaling contributes to abnormal voiding function exhibited by VSA and CMG, we examined SMIR^+/− and SMIR^−/− mice morphologically. As shown in Figure 3H, SMIR^+/− and SMIR^−/− mice bladders are lighter compared with those of wild-type mice. Masson trichrome staining of tissue sections from middle part of bladders from SMIR^+/− and SMIR^−/− mice revealed significantly altered morphology, with bladder wall thickness and muscle layer thickness significantly thinner compared with wild type (Fig. 3). Furthermore, SMIR^+/− and SMIR^−/− bladder muscle bundles are disorganized with enlarged spaces among the bundles, and, particularly in SMIR^−/− bladders, significant interstitial tissue penetrates and segregates the muscle bundles into looser smaller strips (Fig. 3F). The increased proportion of interstitial tissue is obvious in both SMIR^+/− and SMIR^−/− mice bladders, indicating a decompensatory response. Consistent with dilated bladders, the vasculature in SMIR^+/− and SMIR^−/− mice bladder lamina propria layer also dilated (Fig. 3D–F), suggesting a potential significant oxidative stress due to abnormal vascular function, which could contribute to the DBD as well.

Our morphological evidence suggested that IR deletion could impact BSM contractility, thereby leading to altered bladder voiding function. For determination of whether this was true, myography was performed on isolated BSM strips. As shown in Figure 4A–D, the contractile force of BSM strips increases in response to increased EFS frequencies, which mimics the in vivo BSM contraction in response to neurotransmitter release. In comparisons with wild-type mice, the contraction force from SMIR^+/− and SMIR^−/− mice BSM strips was significantly smaller. For confirmation that the diminished contraction force is due to BSM itself, and not due to altered neurotransmitter release, KCl, which depolarizes BSM membrane potential and therefore causes BSM contraction, was used to test BSM response. Results consistently indicated significantly smaller contraction force in BSM strips from SMIR^+/− and SMIR^−/− bladders (Fig. 4E). As noted, bladder contraction is mainly mediated by parasympathetic ATP and Ach co-release, which induce P2x1 receptor– and Chrm3 receptor–mediated signaling cascades leading to BSM contraction. For determination of changes of these signaling pathways in IR-deleted BSM, α,β-meATP, an agonist for P2x1 receptor, and carbachol, an agonist for Chrm3 receptor, were tested, and our data showed significantly impaired contractile force in response to both agonists, suggesting significant change of these pathways in SMIR^+/− and SMIR^−/− mice bladders (Fig. 4F and G).

To further understand the mechanism of impaired smooth muscle contractile function in SMIR^+/− and SMIR^−/− mice, we performed Western blot studies on potential pathways for BSM contraction machinery. Consistent with our functional data showing diminished BSM contraction force (Fig. 4), the expression levels of both Chrm3 receptor and P2x1 receptor were significantly downregulated (Fig. 5A–C). Interestingly, integrin β1, which we have previously shown is an important cell adhesion protein mediating BSM mechanotransduction and voiding function (34), remains unchanged when BSM is IR-null (Fig. 5A and D). However, sm22α, a smooth muscle–specific protein involved in cell differentiation and cytoskeletal organization, is significantly reduced in both SMIR^+/− and SMIR^−/− bladders, suggesting reduced BSM cell proliferation/differentiation. Cav1.2, a voltage-gated ion channel mediating calcium influx, is an essential protein for BSM cell contractile function and gene regulation, and smooth muscle null for Cav1.2 causes a lethal phenotype (33,35). Interestingly, in our SMIR^+/− and SMIR^−/− mice, Cav1.2 is significantly downregulated (Fig. 5A and F), which might also contribute to the diminished contraction force and altered voiding phenotype in these mice.

As noted, insulin signaling is crucial for glucose metabolism, protein/lipid synthesis, and cell proliferation/differentiation in regulating gene transcription. Insulin signaling is mediated by insulin receptor (IR), which belongs to receptor tyrosine kinase (RTK) family and contains the intrinsic tyrosine kinase domain. On activation, IR phosphorylates multiple substrate proteins and modulates many downstream signal pathways. Among them, Akt/protein kinase B, a serine/threonine protein kinase, plays crucial roles by inhibiting Forkhead Box O (FoxO)-mediated protein degradation, autophagy, and apoptosis or by promoting target of rapamycin (mTOR)-mediated metabolism, protein synthesis, and cell proliferation. As expected, IR expression levels in SMIR^+/− and SMIR^−/− mice bladders are significantly reduced (Fig. 6A and B), and the remaining messenger and protein signaling reflects the IR expression from other cell types in bladder wall such as urothelial and interstitial cells. To our surprise, Akt and p-Akt (Akt phosphorylated at Ser473) levels were not changed (Fig. 6A, C, and D), and this could be due to multiple factors. (See discussion.) Interestingly, multiple FoxO proteins are upregulated in SMIR^+/− and SMIR^−/− mice bladders, including FoxO1, FoxO3, and FoxO6 (Fig. 6A and E–G), and correspondingly, mTOR and p-mTOR (mTOR phosphorylated at Ser2448) are significantly decreased in SMIR^+/− and SMIR^−/− mice bladders (Fig. 6A, H, and I). These data indicate an increased catabolism-like protein degradation and autophagy in these mice, which is consistent with our morphological (Fig. 2), functional (Figs. 2 and 4), and molecular (Fig. 5) data shown above.

Adiponectin is a soluble protein secreted mainly by adipocytes, and recent studies have shown that adiponectin-mediated signaling is important in regulating insulin sensitivity, glucose uptake, and lipid metabolism (36,37). Insulin signaling cross talks with adiponectin signaling by interacting with adaptor proteins such as APPL1, and knockdown of IR in adipocytes has been shown to suppress adiponectin production (38). Interestingly, recent studies indicated that vascular smooth muscle cells (VSMC) also express adiponectin, and adiponectin-mediated signaling in VSMC regulates VSMC differentiation and contractile function (39,40). Using immunofluorescent staining and imaging, we determined that adiponectin is also expressed in BSM and in vasculature elements in bladder wall (Fig. 7A and Supplementary Fig. 2). Further Western blot studies indicate that adiponectin protein levels are significantly downregulated in SMIR^+/− and SMIR^−/− mice bladders, suggesting a role of adiponectin signaling in BSM function (Fig. 7B and C). AMPK is a key downstream molecule mediating adiponectin signaling. AMPK plays an important role in cellular energy homeostasis by regulating glucose and fatty acid uptake and oxidation. While there were no detectable changes in AMPK expression levels in SMIR^+/− and SMIR^−/− bladders, p-AMPK (phosphorylated AMPK-α subunit Thr172) levels are inhibited (Fig. 7B, D, and E), suggesting that downregulated adiponectin resulted in reduced AMPK phosphorylation as a consequence of IR deletion.

Insulin signaling is crucial for metabolic homeostasis, and tissue-specific deletion of IR can prompt distinct metabolic disorders (9–11). In a recent report, feeding a high-fat diet for 8 weeks to smooth muscle–specific IR-deleted mice induced obesity in both wild-type and SMIR^−/− mice to reach body weights of ∼40 g. In that case, there was not a significant difference detected between wild-type and SMIR^−/− mice with regard to GTT, ITT, and triglyceride cholesterol levels. The plasma insulin level was somewhat higher in SMIR^−/− mice but did not reach significance (28). However, in that study they did not report on the metabolic condition of SMIR^−/− mice fed on a normal diet. It is possible the high-fat diet could mask abnormal metabolic conditions in these mice. In this study, SMIR^+/− and SMIR^−/− mice fed regular food were smaller compared with wild-type control mice (mostly ranging from 18 to 30 g), and our results indicate that SMIR^−/− mice exhibit normal blood glucose but have impaired GTT and ITT responses (Fig. 1). These data suggest that smooth muscle tissue could contribute to overall metabolic syndromes during stressed condition due to local consumption and storage of glucose.

DBD is an extremely common complication in human patients with diabetes, and they can present with a spectrum of symptoms and urodynamics due to different diabetic cystopathies, including neuronal impairment, BSM damage and areflexia, and urothelial dysfunction (15,41,42). Neuronal impairment and urothelial dysfunction might cause sensory abnormalities resulting in insensate bladder, which leads to increased bladder capacity and overdistended bladder. BSM damage and areflexia might cause diminished BSM contractility and increased postvoid residual volume (PVR). Therefore, patients with diabetes often manifest with urinary frequency, urgency, incontinence, hesitancy, dribbling and weak stream, nocturia, and urinary retention. In our SMIR^+/− and SMIR^−/− mice, we were able to recapitulate many of the symptoms and urodynamic changes observed in human DBD patients, such as frequency (Fig. 2), diminished BSM contractility (Fig. 4), dilated bladder indicating increased bladder capacity (Fig. 3), and small voids (Fig. 2) with dilated bladder (Fig. 3) suggesting urinary retention and increased PVR. However, our conventional CMG study did not generate PVR data in these SMIR^+/− and SMIR^−/− mice, which needs to be further investigated in future with other novel methods such as X-ray videocystometry (43).

Hyperglycemia is often blamed for DBD pathogenesis and polyuria (27). It is believed that hyperglycemia results in oxidative stress and causes cellular damage, and polyuria leads to diuresis and bladder remodeling. In our study, blood glucose levels in both SMIR^+/− and SMIR^−/− mice were normal, and only SMIR^−/− mice exhibited abnormal GTT and ITT responses (Fig. 1 and Supplementary Fig. 1); however, as shown above, significant voiding dysfunctions and morphological changes mimicking human DBD were observed in both homozygous and heterozygous models. This observation implies several important ideas with regard to the pathogenesis of DBD: 1) while hyperglycemia might play some role in the pathogenesis of DBD, it may not be the most essential or major contributing factor. 2) Instead, IR-mediated signaling in bladder itself, which not only regulates bladder tissue metabolic homeostasis but also modulates tissue phenotype and function through regulation of gene expression, could be the major contributing factor leading to DBD pathogenesis. 3) SMIR^+/− and SMIR^−/− mice do not exhibit hyperglycemia (Fig. 1A and Supplementary Fig. 1A) and diuresis and do not exhibit any polyuria (Fig. 2F), both of which are often invoked as causative for DBD. These findings suggest that a major shift may be required in how we think about the etiology of DBD.

In addition to disrupted homeostasis of energy metabolism, what other underlying mechanism(s) might contribute to DBD? In high-fat-fed SMIR^−/− mice, VSMC proliferation in femoral artery at 7 days after wire injury was decreased by 51% compared with wild-type mice, and the intimal hyperplasia upon wire injury was decreased by 37%, with a much thinner femoral arterial wall thickness exhibited (28). These data are fully consistent with our findings that BSM layer is much thinner in SMIR^+/− and SMIR^−/− mice, and the vasculature in the bladder wall is also dilated like the bladder wall (Fig. 3), indicating decreased proliferation in BSM cells due to disrupted insulin signaling. Furthermore, we showed that FoxO1, FoxO3, and FoxO6 protein levels are significantly upregulated, while mTOR and p-mTOR protein levels are significantly downregulated, in SMIR^+/− and SMIR^−/− mice bladder (Fig. 6). Further study is required regarding the exact role of these proteins in BSM function; however, it is generally believed that an increase of FoxO transcription factors and a corresponding decrease of mTOR favors pathways for cell cycle arrest, protein degradation, inflammation, and apoptosis (44). Therefore, IR deletion not only decreases BSM proliferation but also might increase BSM protein degradation and cell apoptosis and thereby change both BSM phenotype and function. This idea is further supported by decreased expression levels of key functional proteins in IR-deficient BSM, including Chm3, P2x1, Cav1.2, and sm22 (Fig. 5), which eventually leads to diminished BSM contraction force (Fig. 4) and voiding dysfunction (Fig. 2).

While only high-fat-fed SMIR^−/− mice were studied in arterial intimal hyperplasia (28), we included both SMIR^+/− and SMIR^−/− mice in this study because we believe the heterozygotes might better mimic the diminished insulin signaling cascades in BSM cells of type 2 diabetes due to decreased insulin release or insulin resistance. Conversely, SMIR^−/− mice might better mimic type 1 diabetes completely lacking in insulin signaling. Interestingly, both mice exhibit significant DBD phenotypes, with SMIR^−/− mice showing relatively more severe symptoms and molecular changes in most cases, and this is in good agreement with the human type 1 and type 2 diabetes situations.

Hyperglycemia, diminished β-cell insulin release, and insulin resistance are key features in human patients with diabetes. Hyperglycemia is a direct consequence of either diminished β-cell insulin release or insulin resistance, which reflects the disruption of normal hepatic regulation of blood glucose by IR-mediated pathway, and in liver-specific IR knockout mice severe hyperglycemia and insulin resistance have been reported, while in other tissue-specific IR knockout studies, including fat, skeletal muscle, and smooth muscle, hyperglycemia has not been shown. However, abnormal tissue/organ function mimicking patients with diabetes has been consistently reported in these studies. Therefore, it seems rational to conclude that disrupted IR signaling in the bladder tissue of human patients with type 1 diabetes and type 2 diabetes is a major pathological mechanism for the development of DBD.

Despite the interesting and novel findings uncovered in these experiments, there are also a number of emerging questions that need be answered in future studies. For example, insulin-like growth factor 1 (IGF1), a growth hormone with a molecular structure similar to that of insulin, can also activate IR signaling. IGF1 receptor (IGF1R) and IR not only form homodimers by themselves but also can form heterodimers with each other. In femoral artery, IGF1R has been shown to be a negative regulator of insulin signaling (28). Elevated IGF1 signaling is suggested to be important in BSM hypertrophy and bladder remodeling (45,46); however, further investigation is needed for detailed understanding of roles of IR- and IGF1R-mediated signaling in bladder physiology and pathogenesis.

Activation of both IR and IGF1R share many common downstream effectors, such as Akt. How different receptors induce differential effects by modulating a common effector is not clear. To our surprise, we observed that Akt and p-Akt expression levels are not affected in SMIR^+/− and SMIR^−/− bladders; however, the downstream FoxO proteins are elevated (Fig. 6). This could be a confounding effect of IGF1 signaling in BSM, or IR signaling in other tissue in bladder wall as well. In wild-type VSMCs, a dose of insulin activation can boost the p-Akt dramatically, while in SMIR^−/− VSMCs diminished p-Akt signaling has been observed (28).

Further insight is also needed regarding the role of adiponectin in BSM, which has never been explored in DBD. Due to the important role of adiponectin in obesity, glucose, and fat metabolism, and its interaction with insulin signaling, we expect that the determination of adiponectin expression in BSM will lead to more discoveries on the details of adiponectin-mediated DBD pathogenesis.

In summary, we have discovered that the bladder phenotype displayed in SMIR^+/− and SMIR^−/− mice recapitulates symptoms and urodynamics observed in human DBD patients, and disruption of IR-mediated proliferation and protein degradation pathways might be the major mechanisms in the pathogenesis of DBD.

Acknowledgments. The authors thank Dr. C. Ronald Kahn’s Laboratory at Harvard Medical School for providing insulin gene floxed mice and Dr. Anne Brunet’s laboratory at Standard University for providing FoxO6 antibody. The authors also thank Dr. Warren Hill (Beth Israel Deaconess Medical Center) for helpful comments on the manuscript.

Funding. The authors acknowledge funding received from the National Institute on Aging/National Institutes of Health (grant R21AG064633 to W.Y.) and National Institute of Diabetes and Digestive and Kidney Diseases/National Institutes of Health (grant R01DK126674 to W.Y.) and financial support provided by the National Institute of Diabetes and Digestive and Kidney Diseases Diabetic Complications Consortium (DiaComp) (https://www.diacomp.org) (grant DK076169 to W.Y.).

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. H.C. performed metabolic measurements, ELISA, VSA, CMG, myography, histological and immunostaining and imaging, and Western blot and analyzed data. A.W. performed Western blot and immunostaining and imaging. W.Y. performed immunostaining and imaging and analyzed data. M.L.Z. contributed to study conception and interpretation of data. W.Y. conceived and supervised the project. H.C. and W.Y. wrote the manuscript, and all authors critically reviewed the manuscript, discussed idea and results, and contributed to the manuscript. W.Y. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.