Testicular blood supply is altered in the 41,XXY* Klinefelter syndrome mouse model

Hypergonadotropic hypogonadism is a major feature of Klinefelter syndrome (KS), assumed to be caused by testicular hormone resistance. It was previously shown that intratesticular testosterone levels in vivo and Leydig cell function in vitro seem to be normal indicating other functional constraints. We hypothesized that impaired testicular vascularization/blood flow could be a co-factor to the observed hypergonadotropic hypogonadism. We evaluated the testicular vascular system by measuring blood vessel sizes during postnatal development and testis blood flow in adult 41,XXY* mice. Proportional distribution and size of blood vessels were analyzed during testicular development (1, 3, 5, 7, 10, 21 dpp, 15 wpp). While ratios of the vessel/testis area were different at 15 wpp only, a lower number of smaller and mid-sized blood vessels were detected in adult KS mice. For testicular blood flow determination we applied contrast enhanced ultrasound. Floating and reperfusion time for testicular blood flow was increased in 41,XXY* mice (floating: XY* 28.8 ± 1.69 s vs XXY* 44.6 ± 5.6 s, p = 0.0192; reperfusion XY* 19.7 ± 2.8 s vs XXY*: 29.9 ± 6.2 s, p = 0.0134), indicating a diminished blood supply. Our data strengthen the concept that an impaired vascularization either in conjunction or as a result of altered KS testicular architecture contributes to hormone resistance.

Scientific RepoRtS | (2020) 10:14369 | https://doi.org/10.1038/s41598-020-71377-0 www.nature.com/scientificreports/ was further strengthened when intratesticular testosterone (ITT) values were found to be comparable between 41,XX Y * and 40,XY* littermates, a finding which was later confirmed in a KS patient cohort 17,18 . Consequently, although the endocrine feedback regulation is definitely disturbed, impaired T production cannot be the reason for the lowered peripheral serum T levels. Testes of KS patients are phenotypically described as small and firm with volume of approximately 2-2.5 mL vs. the healthy normal range above 12 mL per testis 19,20 which is due to a dramatically altered testicular architecture caused by germ cell loss and Leydig cell hyperplasia. We reported previously that the vascularization of testes in 30 weeks old mice was altered mainly due to a changed blood vessel composition 18 . The fact that Leydig cell function is seemingly normal and the blood vessel composition is changed led us to speculate whether an impaired testicular vascularization could contribute to the phenotype of hypergonadotropic hypogonadism.
To address this hypothesis we utilized our 41,XX Y * mouse model to evaluate proportional vessel distribution during postnatal development of the testis and testicular blood flow in adult mice.

Results
Testicular vascularization during development. Testicular vascularization was quantitatively analyzed using central testicular cross sections which underwent immunohistochemical staining for SMA, detecting both, seminiferous tubule walls by marking myoid peritubular cells as well as blood vessels. Thus this staining allowed to clearly distinguish seminiferous tubules from vessels by structural differences and enabled accurate evaluation.We examined testicular cross sections during postnatal development in neonatal (1 dpp), juvenile (3, 5, 7, 10 dpp), pubertal (14, 21 dpp) and adult (15 wpp) 41, XX Y * males and their corresponding 40 XY* littermates as controls. Areas of the entire testicular cross section and those covered by blood vessels and the ratios thereof were measured (Figs. 1, 2a-c). Testis size (area) steadily increased during development in mice of both karyotypes. Significant differences became evident at day 7, 10 and 14 dpp prior to puberty and at 21 dpp resembling the onset of puberty. Differences in testis area were most prominent at 15 wpp in fully adult mice when full spermatogenesis was ongoing in control mice but lacking in 41,XX Y* mice (Fig. 2a). Testicular blood vessel's Vessel categories for vascularization analyses in 40 XY* and 41,XX Y * mice. Upper panel: testicular cross-sections stained for alpha-smooth muscle actin (SMA) to visualize blood vessels (stars) between the seminiferous tubules (st). Such sections were used for quantitative analyses. Lower panel: PAS-stained testicular cross-sections with vessels colored for illustration of size categories (< 80 µm 2 : green, 80-1,000 µm 2 : yellow, > 1,000 µm 2 g: red). Scale bar equals 200 µm.
Scientific RepoRtS | (2020) 10:14369 | https://doi.org/10.1038/s41598-020-71377-0 www.nature.com/scientificreports/ areas were similar in both karyotypes up to 21 dpp, but highly significant differences were noted for time point 15 wpp for 41,XX Y* mice showing a reduced blood vessel area growth (Fig. 2b). Vascularization degree was normalized for the smaller testicular size of 41,XX Y * mice when calculating the ratios of the blood vessels area divided by absolute testis area analyzed (Fig. 2c). Although the degree of vascularization slightly decreased over the early phases of testicular development, no significant changes were noticed until 21 dpp. At 15 wpp, in young adult mice the ratio was significantly higher in 41, XX Y * testes, although the amount of total blood vessels was smaller. This is likely due to the much bigger testis area in 40,XY* males in which spermatogenesis is running at a maximum (Fig. 2a,c). Testicular vascularization according to blood vessels size categories in adult males. Using a three-category system for blood vessels, we noticed that the areas of the smallest (< 80 µm 2 ) vessels belonging to category 1 and the intermediate (80-1,000 µm 2 ) vessels of category 2 were significantly larger in 40,XY* testes (p = 0.0001 respectively). No significant differences between the karyotypes regarding the area covered by the largest vessels (> 1,000 µm 2 ) representing category 3 could be detected, Fig. 3a). For each size category, the respective blood vessel area/testis area ratio was determined, indicating the testicular degree of vascularization (Fig. 3b). The degree of vascularization regarding the smallest blood vessels did not reveal any differences, whereas the degree of vascularization for category 2 and 3 was significantly increased in 41,XX Y* mice (p = 0.0111). Tak-ing advantage of a technology which enables to visualize and measure blood flow in testis in vivo, we addressed the question whether the noticed changes in the blood vessels architecture in 41,XX Y* mice also effects blood flow in the testis (Fig. 4). Contrast enhanced ultrasound analysis revealed significantly smaller testes in 41,XX Y * In (a-c), medians with interquartile ranges are shown; *p < 0.05; **p < 0.01; ***p < 0.001.  Fig. 4a). For blood flow measurements, the perfusion of the testes was videotaped. One major difference was immediately obvious between both groups (Supplementary Video S1) since floating of testes of 41,XX Y *mice was remarkably delayed (Supplementary Video S1, Fig. 5).

Scientific RepoRtS
Using CEUS we then measured the time needed to float and to reperfuse the testes. Time to peak of floating (p = 0.0192) and reperfusion of testes was significantly extended in XX Y * compared to XY* mice. Floating and reperfusion time for testicular blood flow was significantly increased in 41,XX Y * mice (floating: XY* 28.8 ± 1.69 s vs XX Y * 44.6 + 5.6 p = 0.0192; reperfusion XY* 19.7 ± 2.8 s vs XX Y *: 29.9 + 6.2p = 0.0134), indicating a diminished capacity (Figs. 4b, 5a-c, Supplementary Video S1).

Discussion
By using quantitative methods and sophisticated in vivo measurements of the blood flow, this study addresses a mechanism potentially underlying the testicular hormone resistance, which resembles a hallmark of the KS phenotype. We recently reported that LC function and ITT levels are nearly in the normal range in testes of KS patients and in 41,XX Y *mice giving room for the idea that a Leydig cell impairment is not causative for the observed low peripheral T levels. Thus, a different mechanism must be acting on the disturbed endocrine feedback.
When analyzing the testicular vascularization during development, our finding that at 15 weeks of age the area covered by blood vessels related to the entire testis area was significantly larger in 41,XX Y * mice seemingly contradicts our previous observations from 30 week old animals 18 . In mice of this age, we found that the absolute area covered by blood vessels but also the blood vessel to testes area ratio were smaller. This observation might point to a progressive degenerative process, in which the depletion of germ cells, which is complete during the first postnatal weeks 17,21 , affects testicular vascularization in the long-term. The nearly complete lack of germ cells leads to compaction of the testis, which eventually evokes compensatory processes (i.e. additional bigger vessels are formed) possibly responding to the emptied spermatogonial niches. An alternative explanation could be that vessels are still there (resulting in a higher vessel/testis ratio because of the relatively smaller testis size) for a while but get increasingly lost because of the changing testicular architecture, a phenomenon which is also seen in other mouse models losing the germ line 22 . The fact that we see these changes only at 30-weekold animals points to an age-dependent process. It might also well be that at a more advanced age hyalinization of tubules and tubular ghosts, well known to be present in testis of KS patients, are the result of these vascular changes.
Using the CEUS system, we were able to monitor blood flow in vivo to get deeper insights on the effects of the changed vascularization. As we found a significantly increased time to peak values for both, testicular floating as well as reperfusion, we proved that the observed altered vascular situation results in significantly hampered blood flow in the XX Y * testes. It is tempting to assume that the impaired perfusion we observed in 41,XX Y * males is caused by the lack of smaller vessels which prevent a proper distribution of blood within the testis. This phenomenon is prone to worsen at a higher age when the larger vessels are reduced in addition 18 . It might also well be, that progressive testis degeneration and tubular hyalinization are additionally enhanced by a decreasing blood supply, i.e. that a "vicious circle" accelerates the damage of the testicular structures with time, an observation also made in patients with KS 23 .  www.nature.com/scientificreports/ The disturbed testicular circulation may be one co-factor to the endocrine phenotype associated with KS. Only little is known on a putative transport system, which enables gonadotropins, which represent large protein hormones, to pass the blood/testis barrier to interact with their target cell, namely the Sertoli and Leydig cells. It appears plausible that the loss of capillaries in 41,XX Y * mice interferes with this and results in an insufficient stimulation of LC in the presence of highly elevated LH serum levels. In line with this are studies from Belli et al. 24 who further stimulated KS patients with human Chorionic Gonadotropin (hCG) and could evoke an increase in T biosynthesis. However, alternatively, also T release could be hampered, which is supported by the notion that T is produced in sufficient amounts in the testis of KS patients 18 .
Future approaches might prove or disprove this concept experimentally by testing the release of T from testis organoids closely resembling the in vivo situation to elucidate whether testis compactness plays a role. Alternatively, LH function could be studied, e.g. through the administration of small diffusible LHR agonists 24,25 to address the accessibility of LH to the Leydig cells. This approach could clarify the issue to which extent LH action can be significantly increased in the testis.
One of the most prominent and best known features of KS is the progressive loss of germ cells during development, likely to start already pre-to perinatally 21,26 . As a previous study indicated that the localization of SSCs and interstitial cells is related to the testicular vasculature and proposed that the formation of the vascular network precedes and impacts germ line niche establishment 27 , altered vascularization might be involved in these processes or at least they are linked to each other. Two scenarios seem conceivable: the predisposed angiogenetic reduction of the testicular vascularization might provoke the loss of germ cells. Another potential explanation is that the germ cells are depleted due to further undefined causes resulting in an altered testicular architecture that induces a secondary reduction of vascularization. Which of both hypotheses-if any-reflects the actual situation, needs to be investigated in more detail in future studies. In both cases germ cell loss and altered testicular vasculature would impact each other, possibly elucidating a common cause for altered endocrine regulation and germ cell depletion in KS.
It remains to be solved whether X-chromosomal genes escaping from X-inactivation are involved in one or both processes and whether these changes are a secondary consequence of other features of this chromosomal disease. The gene KDM6A/Kdm6a, which escapes X chromosomal inactivation in both, human and this mouse www.nature.com/scientificreports/ model has been shown previously, is known to play a "master switch" role in early organogenesis of the cardiovascular system suggesting a genetic influence on the changes observed in the vascular system in KS 28,29 . Our findings were obtained in a mouse model and although these findings appear to offer a plausible explanation for the hypogonadal status of KS, a confirmation that this observation also holds true in patients remains to be shown. If so, therapeutic approaches aiming at improving testicular vascularization are envisaged.
Taken together, our results for the vasculature architecture in the testes of male 41,XX Y * mice point to a grossly changed vessel composition, including an altered contribution of vessels from the different categories. This ultimately leads to a changed testicular blood flow. Both factors together might play a role for the well-known testicular hormone resistance and germ cell loss in KS.

Materials and methods
Animals. Male 41,XX Y * mice as well as their 40,XY* littermate controls, were obtained from the institutes breeding colony of the B6Ei.Lt-Y* strain 26 . Animals were kept in macrolon type II cages on a layer of wood shavings. The cages were enriched with paper towels as well as red macrolon nest boxes. Animals had ad libitum access to pelletized food (Altromin 1324, Altromin, Lage, Germany) and tap water. Further keeping conditions were a 12:12 LD cycle at a constant room temperature of 24 °C and a humidity of 50%. All applications and protocols were performed in accordance with the national and European legislation for animal care and experiments and were approved by the animal ethics committee of the Landesamt für Natur, Umwelt und Verbraucherschutz Northrhine Wesfalia (regional authorithy for animal ethics LANUV NRW; License numbers: 84-02.05.20.13.115 and 84-02.05.30.12.084). Karyotypes were determined along two different routes using established in house methods. The karyotype of immature mice was determined by Xist-RT-PCR according to a protocol from Werler et al. 30 . Briefly, RNA was isolated from fresh liver tissue by Ultraspec (AMS Biotechnology, Wiesbaden, Germany). RNA (1 lg) was transcribed into cDNA using random Hexamer Primer and Reverse Transcriptase Superscript II (Invitrogen, Darmstadt, Germany). The 83-bp-long Xist fragment was amplified by reverse transcriptase PCR with 1.5 lL cDNA (mouse primer: forward: 5¢-GAG CCC AAA GGG ACA AAC AA-3¢; reverse: 5¢AGT TCT GCT GAG ATG TAA T-GTA GCT GTA TAG -3¢; thermocycling conditions: 3 min 94 °C, 28 cycles: 94 °C: 15 s, 60 °C: 30 s, 72 °C: 1 min; final extension: at 72 °C, 5 min). In contrast, adult mice were karyotyped for presence of sex chromosomes using Fluorescence in Situ Hybridisation of 200-300 µl blood samples. In short, 750 µl Biocoll (Biochrom AG, Berlin, Germany) were used for separation before cells were cracked with KCl and methanol/glacial acetic acid. Of every sample, 0.5 µl was dropped onto a slide and dried on a heating plate. The slides were denatured in 70% formamide 2 × SSC (saline sodium citrate buffer) at 68 °C and dehydrated through an ethanol series. Probes (1200 XMCY3-02; 1189-YMF-02; Cambio, Cambridge, UK) were incubated overnight. Afterwards the slides were placed in 50% formamide 2 × SSC at 43 °C for 15 min, before being transferred into heated 2 × SSC for 10 min and then for another 10 min at room temperature. The slides were counterstained with Hoechst 33258 and mounted in Vectashield (Vector Laboratories, Burlingame, USA) 17,31 . Numbers of animals used in the single experiments are reported in the respective paragraphs.

Contrast enhanced ultrasound analysis (CEUS).
Mice (XX Y *: n = 10, XY* n = 12) aged 15-16 weeks were initially anaesthetized by inhalation of 4.5% isoflurane/l O 2 in a narcosis box (Landmark VSA-2100, Vetland Medical, Louisville, USA) before 1.5% isoflurane/l O 2 was applied continuously during the measurements inside the Vevo imaging station (VisualSonics Inc., Toronto, Canada). Non-targeted contrast agent Vevo MicroMarker™ (VisulaSonics Inc.) solved isotonic sodium chloride (0.9%, B. Braun, Melsungen, Germany) was injected via the tail vein using a catheter (Vevo ® MicroMarker™ TVA Cannulation Kit, VisualSonics) and an ointment (Bepanthen ® , Bayer AG, Leverkusen, Germany) was applied to prevent eye dehydration. The animal was fixed onto the handling table connecting the paws to electrodes of the monitoring unit using contact gel (Aquasonic ® 100, Parker Laboratories, Inc., Fairfield, USA). Animal's heart rate, an electrocardiogram (ECG), body temperature, respiration rate and blood pressure were monitored. The testicular region was shaved to avoid confounding signals, positioned by an elastic strap 32,33 , and de-gassed ultrasound gel (Aquasonic ® 100, Parker Laboratories, Inc., Fairfield, USA) was applied to the scrotum. The Micro Scan™ transducer (MS-250, Visula-Sonics Inc.) was positioned in B-Mode allowing simultaneous observation of both testes with a focus depth of 7-8 mm. Analysis was performed in a contrast specific imaging mode in a two dimensional manner 32,34 . The ultrasound frequency was set to 18 MHz and applied for imaging at 10% of the maximum transmission power and for signal acquisition contrast gain was set to 32 dB to maintain a constant background signal in all measurements. Confounding by respiration was excluded by gating the acquisition between two respiration peaks. Recording started before injection of 45 µl of the agent (rate 15 µl/s) in order to avoid microbubble destruction. When both testes were filled with the contrast agent for the first time, duration of this process was measured; designated "floating". After floating, the microbubbles present in the region of interest [ROI defined by tracing the ultrasound image; the area (mm 2 )] were destroyed by a single ultrasound pulse ("boost"), before testicular reperfusion by the circulating agent was determined, a process designated "reperfusion". The period required ('time to peak') for reaching maximal signal intensity (maximum blood amount in the testis) served as a measure of testicular blood supply for floating and reperfusion.
Histology. All adult animals were sacrificed by cervical dislocation before anaesthesia abated whilst immature animals were killed by immediate decapitation. Both testes were removed and fixed in Bouin's solution for up to 24 h before being dehydrated, and embedded in paraffin. Multiple 5-µm sections of each sample were obtained. Every second slide was stained with periodic acid-Schiff (PAS)/hematoxylin 17  www.nature.com/scientificreports/ central testicular part of adults was processed, serial sections generated, and the section with greatest diameter selected for subsequent evaluation of vascularization by immunostaining against a-smooth muscle actin (SMA, see below).

Analysis of vascularization.
The vascular composition in testicular tissues of 41,XX Y* and 40,XY* littermate mice was histologically evaluated at 1 dpp (XX Y *: n = 5, XY* n = 7), 3 dpp (XX Y *: n = 5, XY* n = 5), 5 dpp (XX Y *: n = 5, XY* n = 6), 7 dpp (XX Y *: n = 5, XY* n = 6), 10 dpp (XX Y *: n = 5, XY* n = 5), 14 dpp (XX Y *: n = 5, XY* n = 5), 21 dpp (XX Y *: n = 5, XY* n = 6) dpp and 15-16 (XX Y *: n = 10, XY* n = 12) wpp. Developmental stages were chosen in consistence with previously published studies 17,21 . PAS stained cross sections taken from the center of each testis were analyzed microscopically for identification of the testicular region with the greatest circumference. Corresponding slides from this central region were then immunohistochemically stained against SMA for detection of the vessels and serial pictures covering the entire area were taken or, if required, single pictures were assembled and analyzed as previously published (magnification 100×; Olympus BX61, Adobe Photoshop CS3, Version 10.0 (Adobe Systems Incorporated, San Jose, USA 18 ). To address testicular blood vessel composition during development we assigned the vessels to three categories. (1) smaller than 80 µm 2 , which resembles mainly capillaries; (2) larger than 80 µm 2 but smaller than 1,000 µm 2 indicative for intermediate and (3) larger than 1,000 µm 2 for bigger venules and arterioles (Fig. 1). From the region of testis with the greatest diameter two sections per animal were evaluated in all developmental stages. Per developmental stage and karyotype at least 200 vessels were evaluated ranging from 32 to 102 vessels per animal.
Immunohistochemical staining (IHC) for alpha smooth actin (SMA). For detection and size measurement of blood vessels, IHC against SMA was performed according to a protocol previously published 35 . We used a primary antibody directed against smooth muscle actin (SMA; mouse-monoclonal anti-SMA, A2547, 1:1000, Sigma-Aldrich, Hamburg, Germany) which is encoded by the ACTA2 gene. Testicular sections treated with corresponding mouse (I5381) IgG antibodies served as control (1:1000, Sigma-Aldrich, Hamburg, Germany). After overnight incubation, sections were washed three times in TBS and the corresponding secondary antibody, conjugated to horse-radish peroxidase (chicken anti-mouse HRP, sc2954, 1:100, Santa Cruz Biotechnology, Inc. Heidelberg, Germany) were applied. Protein expression was visualized using 3,3′-diaminobenzidine as chromogen. Hematoxylin was applied as counterstain (Fig. 1). Samples were analyzed microscopically and documented as described above.
Serum testosterone determination. Trunk blood was collected and measurement of serum T concentrations in adult (15 week) 40,XY and 41,XX Y * male mice from the enhanced ultrasound experiments were conducted utilizing a solid-phase double-antibody radioimmunoassay (RIA). The RIA was performed using iodinated tracer (testosterone-3-(O-carboxymethyl) oximino-2-[125I] iodohistamine, Amersham International, Germany), a rabbit testosterone-3 (carboxymethyloxime)-BSA antiserum and a secondary antibody against rabbit IgG (Bio-Rad Laboratories Incorporation, Hercules, USA). Recovery after ether extraction was monitored by addition of tracing amounts of [1β, 2β-3H] testosterone (NEN Life Sciences, Inc., Boston, MA, USA) to the sera and the results were corrected accordingly. Detection limit was 0.72 nmol/l. Duplicate serum samples from each animal were analyzed for serum T levels. The intra-and inter-assay coefficients of variation were 4.7 and 5.8%, respectively 36-38 . Statistics. Statistical analysis was performed using Microsoft Excel 2010 (Microsoft Corporation, Redmond, USA) and GraphPad Prism Version 5.0 (GraphPad Software, San Diego, USA). Due to group sizes and as missing normal distribution the nonparametric Mann Whitney U test was utilized for data analysis. For testosterone measurements, due to the small sample numbers, a Gaussian distribution could not be assumed and a regular two-way ANOVA and Bonferroni posttests were performed. Values were considered significantly different when p < 0.05. If not stated otherwise, medians are given.