Abstract
Spermatogenesis, the complex developmental process of male germ cell proliferation, differentiation, and maturation, is the basis of male fertility and reproductive fitness. In the seminiferous tubules of the testes, spermatozoa are constantly generated from spermatogonial stem cells through a stereotyped sequence of mitotic and meiotic divisions. The basic physiological principles, however, that control both maturation and luminal transport of the still immotile spermatozoa within the seminiferous tubules remain poorly, if at all, defined. Here, we show that coordinated contractions of smooth muscle-like testicular peritubular cells provide the propulsive force for luminal sperm transport towards the rete testis and epididymis. Using a mouse model for in vivo imaging, we describe and quantify spontaneous tubular contractions and show a causal relationship between peritubular Ca2+ waves and peristaltic transport. Moreover, we identify P2 receptor-dependent purinergic signaling pathways as physiological triggers of tubular contractions both in vitro and in vivo. When challenged with extracellular ATP, transport of luminal content inside the seminiferous tubules displays stage-dependent directionality. We thus suggest that paracrine purinergic signaling coordinates peristaltic recurrent contractions of the mouse seminiferous tubules to propel immotile spermatozoa to the rete testis. Consequently, our findings could have substantial pharmaceutical implications for both infertility treatment and / or male contraception.
Introduction
Spermatogenesis ranks among the most complex, yet least understood developmental processes in postnatal life. Inside the seminiferous tubules, this intricate course of mass cell proliferation and transformation events generates haploid spermatozoa from diploid spermatogonial stem cells. Along the seminiferous epithelium, spermatogenesis has been conceptualized by attribution of sequential cellular ‘stages’, which progress through coordinated cycles (Russell, 1990). Each spermatogenic cycle completes with the release of immotile spermatozoa from the seminiferous epithelium into the lumen of the tubule. Once detached from the Sertoli cells, sperm must be transported to the rete testis and epididymis for final maturation. Precisely regulated tubular transport mechanisms are, thus, imperative for reproduction.
While bulk movement of luminal content has been anecdotally reported (Cross, 1958; Setchell et al., 1978; Worley et al., 1985), no quantitative data on sperm transport within the seminiferous tubules is available. Early in vitro observations of apparent minute undulating motions of seminiferous tubule segments (Roosen-Runge, 1951; Suvanto and Kormano, 1970) suggested that smooth muscle-like testicular peritubular cells (TPCs) (Clermont, 1958; Ross, 1967) could mediate contractile tubule movements. This concept has gained widespread support from several, mostly indirect, in vitro studies (Ailenberg et al., 1990; Filippini et al., 1993; Miyake et al., 1986; Tripiciano et al., 1996). However, quantitative direct (i.e., live cell) measurements of seminiferous tubule contractions are rare and controversial (Ellis et al., 1978; Harris and Nicholson, 1998; Losinno et al., 2012; Worley and Leendertz, 1988). Moreover, mechanistic in vivo evidence is lacking. Here, we demonstrate that, by acting on ionotropic and metabotropic P2 receptors, extracellular ATP activates TPC contractions that trigger directional sperm movement within the seminiferous tubules both in vitro and in vivo.
Results
ATP is a potent TPC stimulus
Accumulating data suggests that purinergic signaling constitutes a critical component of testicular paracrine communication (Fleck et al., 2016; Foresta et al., 1995; Gelain et al., 2003; Poletto Chaves et al., 2006; Veitinger et al., 2011a; Walenta et al., 2018), with Sertoli cells acting as a primary source of ATP secretion (Gelain et al., 2005). Therefore, we asked if mouse TPCs are sensitive to extracellular ATP. Primary TPC cultures retain high purity for ≥14 days in vitro (Figure 1A, and Figure 1–figure supplement 1A&B) and cells express transcripts for several ionotropic (P2X2, P2X4, P2X7) and metabotropic (P2Y2, P2Y6) purinoceptors (Figure 1B). The specific biophysical and pharmacological profile of ATP-dependent transmembrane currents – i.e., saturation at ≤100 μM ATP, modest but persistent desensitization, reduced BzATP sensitivity (data not shown), potentiation by ivermectin, and inhibition by suramin (Figure 1C-G) – strongly suggests functional expression of P2X2 and / or P2X4, but not P2X7 receptors (Coddou et al., 2011). Notably, ~46% of all ATP-sensitive TPCs additionally displayed a delayed, but long-lasting inward current that gradually developed over tens of seconds after ATP stimulation ended (Figure 1–figure supplement 1C&D). Also activated by UTP, dependent on presence of intracellular GTP, and largely carried by Cl− (Figure 1–figure supplement 1E-H), this current likely results from P2Y receptor-mediated phosphoinositide turnover, Ca2+ release, and activation of Ca2+-gated Cl− channels. Notably, live-cell ratiometric Ca2+ imaging in TPCs revealed robust and repetitive cytosolic Ca2+ transients upon ATP exposure (Figure 1H&I). Reducing extracellular Ca2+ to 100 nM, a concentration approximately equimolar to cytosolic levels, did substantially reduce, but not abolish ATP response amplitudes (Figure 1J&K). The selective P2Y receptor agonist UTP also triggered Ca2+ signals (data not shown). Together, these data suggest that mouse TPCs functionally express both ionotropic and metabotropic purinoceptors.
Next, we asked if TPCs also exhibit ATP sensitivity in their physiological setting. In parallel approaches, we examined purinergic Ca2+ signals from mouse TPCs in acute seminiferous tubule sections (Fleck et al., 2016), using either TPC-specific expression of genetically encoded Ca2+ indicators (GCaMP6f) or bulk loading with a synthetic Ca2+ sensor (fura-2). First, we confirmed inducible smooth muscle-targeted testicular expression of fluorescent reporter proteins in SMMHC-CreERT2 x Ai14D mice (Figure 2A–C, movie S1). Second, TPC-specific GCaMP6f expression in SMMHC-CreERT2 x Ai95D mice revealed robust Ca2+ transients in cells of the tubular wall upon ATP exposure (Figure 2D&E). Third, fura-2/AM loading of intact seminiferous tubule sections preferentially labelled the outermost cell layer (Figure 2F), allowing semi-quantitative in situ imaging of ATP-dependent Ca2+ signals in mouse TPCs (Figure 2G&H). So far, our results thus demonstrate that challenging TPCs with extracellular ATP triggers robust Ca2+ signals both in vitro and in situ.
ATP triggers seminiferous tubule contractions
We hypothesized that ATP-induced Ca2+ signals in TPCs could mediate contractile motion of the seminiferous tubule. To address this, we established a fast, quasi-simultaneous image acquisition method that enables parallel recording of both peritubular Ca2+ responses and seminiferous tubule movement (methods). Brief ATP exposure resulted in a peripheral band of Ca2+ activity at the edge of the tubule. Such signals usually coincided with a pronounced contractile motion of the seminiferous tubule (Figure 3A, movie S2). When movement is quantified as the time-lapse image flow field strength (methods) tubular contraction follows the Ca2+ signal onset with minimal delay, outlasts the Ca2+ signal peak, and recovers slowly (Figure 3B). Both Ca2+ responses and tubular movement are dose-dependent and share an ATP threshold concentration of approximately 1 μM (Figure 3C-E, movie S3). Notably, in some tubules, we observed spontaneous low-amplitude ‘vibratory’ movements and local indentations (Figure 4A), reminiscent of the relatively high frequency rippling previously described (Ellis et al., 1981; Worley et al., 1985).
We next investigated the Ca2+ signaling mechanism(s) underlying ATP-dependent TPC contractions. First, we asked whether influx of external Ca2+ is involved in TPC force generation. Similar to in vitro observations (Figure 1J&K), diminishing or even reversing the driving force for transmembrane Ca2+ flux by reducing the extracellular Ca2+ concentration to 100 nM or 12 nM, respectively, for variable durations, significantly decreased both TPC Ca2+ signals and tubular contractions (Figure 4B-H). These effects were fully reversible. Second, we examined a potential role of ATP-induced Ca2+ release from internal storage organelles. Ca2+ depletion of the sarcoplasmic reticulum via pharmacological inhibition of the sarco / endoplasmic reticulum Ca2+-ATPase essentially abolished both ATP-dependent Ca2+ signals and contractions (Figure 4D). Importantly, all results from ratiometric fura-2 imaging were qualitatively indistinguishable from those obtained with genetically targeted GCaMP6f (Figure 4E-G). Third, we aimed to quantify the specific contribution of metabotropic purinoceptors to the overall ATP-mediated effect. The P2Y receptor-selective agonist UTP evoked both TPC Ca2+ signals and tubular contractions (Figure 4E&F). However, these responses were substantially reduced compared to control ATP stimulations (Figure 4F), similar to the diminished signals observed under low extracellular Ca2+ conditions.
Together, these data strongly suggest that (i) extracellular ATP acts as a potent TPC stimulus that triggers seminiferous tubule contractions in situ, that (ii) both P2X and P2Y receptors mediate TPC responses to ATP exposure, that (iii) Ca2+ mobilization from the sarcoplasmic reticulum is necessary to evoke TPC responses, and consequently that (iv) influx of external Ca2+ via ionotropic P2X receptors is not sufficient to drive TPC signals and evoke contractions. Notably, our general finding of ATP-induced mouse TPC contractions is likely transferable to human peritubular cells. When primary human TPC cultures (Walenta et al., 2018) were exposed to extracellular ATP, morphological changes were observed within seconds-to-minutes (Figure 4–figure supplement 1A&B). Moreover, embedding cells in collagen gel lattices revealed considerable contractile force in response to ATP (Figure 4–figure supplement 1C&D).
ATP drives directional luminal transport
We hypothesized that ATP-induced tubular contractions could impact the transport of luminal fluid and spermatozoa. To test this, we custom-built a whole-mount macroscopic imaging platform, designed to allow both widefield and fluorescence time-lapse imaging of intact seminiferous tubules (Figure 5A). In addition, this setup enables visual categorization of the twelve seminiferous epithelium cycle stages into three distinct stage groups (Hess and De Franca, 2008) and allows precisely timed focal perfusion (methods). First, we asked if brief focal purinergic stimulation triggers seminiferous tubule contractions and, consequently, luminal content movement. Flow field change analysis reveals some basal luminal motion independent of mechanical stimulation (Figure 5B). However, ATP exposure triggered a strong increase in luminal flow field strength that outlasted the presence of ATP for several tens of seconds (Figure 5B, movie S4). Second, we analyzed if luminal movement depends on the tubule’s cycle stage and, consequently, luminal sperm count. When we compared ATP-induced movement between directly stimulated stages (i.e., region-of-interest (ROI) 0) with a high (group II) versus a relatively low (groups I & III) amount of luminal sperm, we observed no difference in stimulation-dependent motion (Figure 5CI). Thus, direct ATP exposure triggers tubular contractions independent of cycle stage and luminal sperm count. Third, we investigated if luminal movement is restricted to the area of stimulation or, by contrast, if motion propagates beyond the directly stimulated tubule section. When we analyzed luminal motion in equidistant tubule sections adjacent to the directly stimulated area (Figure 5A), we found a significant, though relatively small bidirectional wave of propagating movement in stage groups with a low luminal sperm count (Figure 5CII). Strikingly, we observed strong unidirectional movement upon ATP stimulation of tubule sections with high luminal sperm density (Figure 5CII). In this stage group, luminal content is predominantly propelled towards areas of ascending spermatogenic cycle stages. These findings demonstrate directionality of sperm transport upon purinergic TPC stimulation.
As expected, ATP-induced tubule contractions also manifest as Ca2+ signals in TPCs (Figure 6A&B, movie S5). Surprisingly, however, these Ca2+ elevations are limited to those areas directly exposed to ATP. We observed no such signals in adjacent tubule sections independent of the stimulated stage group and an ascending or descending stage direction (Figure 6C&D). This finding indicates that ATP acts as a local paracrine messenger that, by itself, is not sufficient to trigger a signal that propagates in a regenerative wave-like fashion along a tubule’s longitudinal axis. In turn, transport directionality likely results from morphological characteristics rather than peristaltic contractility.
ATP induces tubular contractions in vivo
To ultimately attribute a physiological role to ATP-dependent Ca2+ signals in TPCs, tubular contractions, and corresponding transport of luminal content, these phenomena must (i) occur spontaneously in living animals, and must (ii) be triggered experimentally by ATP exposure in vivo. Thus, to investigate any in vivo relevance of our findings, we designed a custom-built 3D printed in vivo imaging stage (Figure 7–figure supplement 1) that allows both widefield epi-fluorescence and multiphoton microscopy of the mouse testis.
Initially, we monitored spontaneous seminiferous tubule activity in SMMHC-CreERT2 x Ai95D mice. Multiphoton time-lapse imaging revealed spontaneous TPC Ca2+ signals that typically accompanied strong tubule contractions (Figure 7A&B, movie S6). Several characteristics emerged from quantitative analysis of these observations. First, during sufficiently long recording periods (≤10 min), contractions occur in essentially all seminiferous tubules (Figure 7–figure supplement 2A). Second, contractions of individual tubules are not synchronized (Figure 7B). Third, periods of enhanced activity (≥2 contractions within 90 s) are interrupted by long episodes of quiescence (Figure 7B, Figure 7–figure supplement 2B). Fourth, the durations of TPC Ca2+ signals and corresponding contractions are positively correlated (Figure 7–figure supplement 2C), confirming a causal relationship.
Next, we asked whether spontaneous in vivo contractions are coordinated along the longitudinal tubular axis. Low magnification incident light microscopy enabled simultaneous observation of several superficial seminiferous tubule segments (Figure 7C). Movement analysis along the length of digitally straightened tubules demonstrates wave-like unidirectional motions that propagate with high velocities (Figure 7C&D). Notably, these coordinated contractile movements provide sufficient force to ensure luminal sperm transport (movie S7).
Finally, we examined if brief focal ATP stimulation also triggers peritubular Ca2+ signals and seminiferous tubule contractions in vivo. Therefore, we filled low resistance patch pipettes with fluorescently labelled ATP solution, penetrated the tunica albuginea, and targeted the interstitial space close to neighboring tubules (Figure 7E, movie S8). Nanoliter puffs of ATP-containing test solution induced both Ca2+ transients in genetically labeled TPCs and strong tubule contractions in the majority of experiments (Figure 7F&G). By contrast, puffs of extracellular saline rarely stimulated any such response (Figure 7–figure supplement 2D). Taken together, in vivo recordings demonstrate that robust recurrent seminiferous tubule contractions (i) occur spontaneously, (ii) are driven by cytosolic Ca2+ elevations in TPCs, and (iii) can be triggered experimentally by ATP exposure. Consequently, paracrine purinergic signaling in the mouse testis is a mediator of luminal sperm transport within the seminiferous tubule network.
Discussion
The molecular and cellular mechanisms that control paracrine testicular communication have to a large extent remained controversial, if not elusive (Schlatt and Ehmcke, 2014). For TPCs in particular, a contractile function under paracrine control and, consequently, a critical role in male infertility have long been proposed (Albrecht et al., 2006; Romano et al., 2005), but direct experimental evidence has been lacking (Mayerhofer, 2013). While several signaling molecules, including vasopressin (Pickering et al., 1989), oxytocin (Worley et al., 1985), prostaglandins (Hargrove et al., 1975), endothelin (Filippini et al., 1993), and others (Albrecht et al., 2006; Mayerhofer, 2013), have been proposed to act on TPCs, a role of ATP in seminiferous tubule contractility has been explicitly ruled out early on (Hovatta, 1972). By contrast, our data reveal ATP is a strong stimulus that activates TPCs via P2X and P2Y receptors, mediating coordinated tubule contractions and luminal sperm transport in situ and in vivo. Both spontaneous and ATP-dependent contractions trigger fast, stage-dependent, and directional transport of luminal content. It is thus tempting to speculate that seminiferous tubule contractility in general, and purinergic TPC signaling in particular, are promising targets for male infertility treatment and / or contraceptive development.
Materials and Methods
Animals
All animal procedures were approved by local authorities and in compliance with both European Union legislation (Directive 2010/63/EU) and recommendations by the Federation of European Laboratory Animal Science Associations (FELASA). When possible, mice were housed in littermate groups of both sexes (room temperature (RT); 12:12 h light-dark cycle; food and water available ad libitum). If not stated otherwise, experiments used adult (>12 weeks) males. Mice were killed by CO2 asphyxiation and decapitation using sharp surgical scissors. We used C57BL/6J mice (Charles River Laboratories, Sulzfeld, Germany) as well as offspring from crossing either SMMHC-CreERT2 (JAX #019079) (Wirth et al., 2008) or 129S.FVB-Tg(Amh-cre)8815Reb/J (JAX #007915) (Holdcraft and Braun, 2004) mice with either Ai95D (JAX #028865) (Madisen et al., 2015) or Ai14D (JAX #007914) (Madisen et al., 2010) mice, respectively.
Chemicals and solutions
The following solutions were used:
(S1) 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES) buffered extracellular solution containing (in mM) 145 NaCl, 5 KCl, 1 CaCl2, 0.5 MgCl2, 10 HEPES; pH = 7.3 (adjusted with NaOH); osmolarity = 300 mOsm (adjusted with glucose).
(S2) Oxygenated (95% O2, 5% CO2) extracellular solution containing (in mM) 120 NaCl, 25 NaHCO3, 5 KCl, 1 CaCl2, 0.5 MgCl2, 5 N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid (BES); pH = 7.3; 300 mOsm (glucose).
(S3) Extracellular low Ca2+ solution containing (in mM) 145 NaCl, 5 KCl, 0.5 MgCl2, 10 HEPES; pH = 7.3 (NaOH); osmolarity = 300 mOsm (glucose); [Ca2+]free = ~110 nM (1 mM EGTA, 0.5 mM CaCl2) or 12 nM (1 mM EGTA, 0.1 mM CaCl2).
(S4) Oxygenated (95% O2, 5% CO2) extracellular solution containing (in mM) 120 NaCl, 25 NaHCO3, 5 KCl, 0.5 MgCl2, 5 BES; pH = 7.3; 300 mOsm (glucose); [Ca2+]free = 110 nM (1 mM EGTA, 0.5 mM CaCl2) or 12 nM (1 mM EGTA, 0.1 mM CaCl2).
(S5) Gluconate-based extracellular solution containing (in mM) 122.4 Na gluconate, 22.6 NaCl, 5 KCl, 1 CaCl2, 0.5 MgCl2, 10 HEPES; pH = 7.3 (adjusted with NaOH); osmolarity = 300 mOsm (glucose).
(S6) Standard pipette solution containing (in mM) 143 KCl, 2 KOH, 1 EGTA, 0.3 CaCl2, 10 HEPES ([Ca2+]free = 110 nM); pH = 7.1 (adjusted with KOH); osmolarity = 290 mOsm (glucose).
(S7) Gluconate-based pipette solution containing (in mM) 110 Cs gluconate, 30 CsCl, 2 CsOH, 1 EGTA, 0.3 CaCl2, 10 HEPES ([Ca2+]free = 110 nM); pH = 7.1 (adjusted with CsOH); osmolarity = 290 mOsm (glucose).
In some experiments Na-GTP (0.5 mM) was added to the pipette solution. Free Ca2+ concentrations were calculated using WEBMAXCLITE v1.15. If not stated otherwise, chemicals were purchased from Sigma (Schnelldorf, Germany). Cyclopiazonic-acid (CPA) and 2’(3’)-O-(4-Benzoylbenzoyl)adenosine-5’-triphosphate (BzATP) triethylammonium salt was purchased from Tocris Bioscience (Bristol, UK). Fura-2/AM was purchased from Thermo Fisher Scientific (Waltham, MA). Final solvent concentrations were ≤0.1%. When high ATP concentrations (≥1 mM) were used, pH was readjusted.
Stimulation
For focal stimulation, solutions and agents were applied from air pressure-driven reservoirs via an 8-in-1 multi-barrel ‘perfusion pencil’ (AutoMate Scientific; Berkeley, CA). Changes in focal superfusion (Veitinger et al., 2011a) were software-controlled and, if required, synchronized with data acquisition by TTL input to 12V DC solenoid valves using a TIB 14S digital output trigger interface (HEKA Elektronik, Lambrecht/Pfalz, Germany). For focal stimulation during in vivo recordings, ATP was puffed from pulled glass pipettes using a microinjection dispense system (Picospritizer III; Parker Hannifin, Hollis, NH).
Extracellular low Ca2+ solutions (S3 & S4) were applied via both the bath and perfusion pencil. To ensure depletion of Ca2+ stores by CPA we monitored intracellular Ca2+ levels during drug treatment (0.05 Hz frame rate). Transient CPA-dependent Ca2+ elevations lasted 10 – 40 min. After baseline Ca2+ levels were restored, cells / slices were again challenged with ATP. Control recordings, omitting CPA, were performed under the same conditions.
Slice preparation
Acute seminiferous tubule slices were prepared as previously described (Fleck et al., 2016) with minor modifications. Briefly, seminiferous tubules from young adults were isolated after tunica albuginea removal, embedded in 4% low-gelling temperature agarose (VWR, Erlangen, Germany), and 250 μm slices were cut with a VT1000S vibratome (Leica Biosystems, Nussloch, Germany). Acute slices were stored in a submerged, oxygenated storage container (S2; RT). When using testicular tissue from Ai95D mice, slices were protected from light during storage to avoid GCamP6f bleaching.
TPC culture
After mouse testis isolation and removal of the tunica albuginea, the seminiferous tubules were placed in Dulbecco’s Modified Eagle Medium / Nutrient Mixture F-12 (DMEM/F-12; Invitrogen) containing 1 mg ml−1 collagenase A and 6 μg ml−1 DNase (10 min; 34°C; shaking water bath (60 cycles min−1)). Three times, the samples were washed (DMEM/F-12; 5 ml), allowed to settle for 5 min, and the supernatant was discarded. Next, tubules were incubated DMEM/F-12 containing 1 mg ml−1 trypsin and 20 μg ml−1 DNase (20 min; 34°C; shaking water bath (60 cycles min−1)). Digestion was stopped by addition of 100 μg ml−1 soybean trypsin inhibitor (SBTI) and 20 μg ml−1 DNase in phosphate-buffered saline (D-PBS). Then, samples were allowed to settle for 5 min and the supernatant was collected. After two more cycles of washing (DMEM/F-12), settling (5 min), and supernatant collection, the collected cell suspension was centrifuged (10 min; 400g) and the supernatant discarded. The pellet was resuspended in DMEM containing FBS (10%) and penicillin G / streptomycin (1%), filtered (cell strainer (100 μm)), and cells were plated in 75 cm2 cell culture flask (T75; Invitrogen) and placed in a humidified incubator (37°C; 5% CO2). Approximately ⅓ of medium volume was replaced every 3 days. Cells usually reached 100% confluence after 7 days in vitro (DIV). Then, cells were washed twice (DPBS−/−; 5 min; 37°C) and incubated in 0.05% trypsin / EDTA (5 min; 37°C). Detachment of cells was checked visually and, if necessary, facilitated mechanically. The cell suspension was centrifuged (3 min; 800g) and the supernatant discarded. The pellet was resuspended in DMEM at cell densities of ~105 cells ml−1 and plated again either in culture flasks or on glass coverslips in 35 mm dishes for experimental use. Again, ⅓ of medium volume was replaced every 3 days. Experiments were performed for ≤5 days after passage.
Human TPCs were isolated from small testicular tissue fragments derived from consenting donors with obstructive azoospermia and normal spermatogenesis as described (Albrecht et al., 2006; Walenta et al., 2018). The study was approved by the local ethical committee (Ethikkommission, School of Medicine, TU Munich, project 169/18S).
Gene expression analysis
Total RNA was isolated and purified from cultured mouse TPCs (passage 1) with Trizol followed by complementary DNA synthesis with RevertAid™ H Minus kit (#K1632 Thermo Fisher) according to the manufacturer’s instructions. Controls in which the reverse transcriptase was omitted were routinely performed. PCR amplification was performed during 30 thermal cycles (95°C, 20 s; 58°C, 20 s; 72°C, 20 s). The following specific primer pairs were used for PCR amplification:
Immunochemistry and tissue clearing
For immunochemistry of testicular cryosections, testes were fixed with 4% (w/v) paraformaldehyde (PFA) in PBS−/−(10 mM, pH 7.4; ≥12 h; 4°C) and subsequently cryoprotected in PBS−/−containing 30% sucrose (≥24 h; 4°C). Samples were then embedded in Tissue Freezing Medium (Leica Biosystems), sectioned at 20 μm on a Leica CM1950 cryostat (Leica Biosystems), and mounted on Superfrost Plus slides (Menzel, Braunschweig, Germany). For immunostaining of cultured mouse TPCs, cells were washed (3x; PBS−/−), fixed with ice-cold 4% PFA in PBS−/−(20 min; RT), and washed again (3x; PBS−/−). For blocking, sections / cells were incubated in PBS−/− containing Tween-20 (0.1%) / BSA (3%) solution (1 h; RT). After washing (PBS−/−; 2 × 5 min), sections / cells were incubated FITC-conjugated monoclonal anti-actin, α-smooth muscle (α-SMA-FITC, cat # F3777, MilliporeSigma) antibody (1:500 in 3% BSA; 1 h; RT). Excess antibodies were removed by washing (2 × 5 min PBS−/−). For nuclear counterstaining, sections / cells were then incubated in PBS−/− containing either DAPI (5 μg ml−1; 10 min; RT; Thermo Fisher Scientific) or DRAQ5 (1:500; 5 min; RT; Thermo Fisher Scientific).
Fluorescent images were taken using either an inverted microscope (Leica DMI4000B, Leica Microsystems) or an upright fixed stage scanning confocal microscope (TCS SP5 DM6000 CFS; Leica Microsystems) equipped with a 20x 1.0 NA water immersion objective (HCX APO L; Leica Microsystems). To control for non-specific staining, experiments in which the primary antibody was omitted were performed in parallel with each procedure. Digital images were uniformly adjusted for brightness and contrast using Adobe Photoshop CS6 (Adobe Systems, San Jose, CA, USA).
For testicular tissue clearing we adopted the CLARITY method (Chung et al., 2013) with minor modifications (Gretenkord et al., 2019). Briefly, testes from adult mice were fixed overnight at 4°C in hydrogel fixation solution containing 4% acrylamide, 0.05% bis-acrylamide, 0.25% VA-044 Initiator, 4% PFA in PBS−/− to maintain structural integrity. After hydrogel polymerization, lipids were removed by incubation in 4% sodium dodecyl phosphate (SDS) solution with 200 mM boric acid (pH 8.5) over periods of two months. Solutions were changed bi-weekly. During the final incubation period, the nuclear marker DRAQ5 (1:1000) was added. After washing (2 d) with PBST (0.1% TritonX), samples were incubated for 24 h in RIMS80 containing 80 g Nycodenz, 20 mM PS, 0.1% Tween 20, and 0.01% sodium acid. Cleared samples were imaged using a Leica TCS SP8 DLS confocal microscope, equipped with a digital light-sheet module, 552 nm and 633 nm diode lasers, a HC PL FLUOTAR 5x/0.15 IMM DLS objective (observation), a L 1.6x/0.05 DLS objective (illumination), a DLS TwinFlect 7.8 mm Gly mirror cap, and a DFC9000 sCMOS camera. Rendering and three-dimensional reconstruction of fluorescence images was performed using Imaris 8 microscopy image analysis software (Bitplane, Zurich, Switzerland).
Electrophysiology
Whole-cell patch-clamp recordings were performed as described (Fleck et al., 2016; Veitinger et al., 2011b). Briefly, mouse TPCs were transferred to the stage of an inverse microscope (DMI 4000B, Leica Microsystems), equipped with phase contrast objectives and a cooled CCD-camera (DFC365FX, Leica Microsystems). Cells were continuously superfused with solution S1 (~3 ml min−1; gravity flow; ~23°C). Patch pipettes (~5 MΩ) were pulled from borosilicate glass capillaries with filament (1.50 mm OD / 0.86 mm ID; Science Products) on a PC-10 vertical two-step micropipette puller (Narishige Instruments, Tokyo, Japan), fire-polished (MF-830 Microforge; Narishige Instruments) and filled with S6. An agar bridge (150 mM KCl) connected reference electrode and bath solution. An EPC-10 amplifier controlled by Patchmaster 2.9 software (HEKA Elektronik) was used for data acquisition. We monitored and compensated pipette and membrane capacitance (Cmem) as well as series resistance (Rseries). Cmem values served as a proxy for the cell surface area and, thus, for normalization of current amplitudes (i.e., current density). Cells displaying unstable Rseries values were not considered for further analysis. Liquid junction potentials were calculated using JPCalcW software (Barry, 1994) and corrected online. Signals were low-pass filtered [analog 3- and 4-pole Bessel filters (−3 dB); adjusted to ⅓ - ⅕ of the sampling rate (10 kHz)]. Holding potential (Vhold) was −60 mV.
Fluorescence Ca2+ imaging
Cultured mouse TPCs were imaged as described (Veitinger et al., 2011b). Briefly, cells were loaded with fura-2/AM in the dark (5 μM; 30 min; RT; S1) and imaged with an upright microscope (Leica DMI6000FS, Leica Microsystems) equipped for ratiometric live-cell imaging with a 150W xenon arc lamp, a motorized fast-change filter wheel illumination system for multi-wavelength excitation, a CCD camera (DFC365 FX, Leica), and Leica LAS X imaging software. Ten to thirty cells in randomly selected fields of view were viewed at 20x magnification and illuminated sequentially at 340 nm and 380 nm (cycle time 2 s). The average pixel intensity at 510 nm emission within user-selected ROIs was digitized and calculated as the f340/f380 intensity ratio.
For parallel recordings of intracellular Ca2+ signals and tubular contractions, acute seminiferous tubule slices were bulk-loaded with fura-2/AM in the dark (30 μM; 30 min; RT). After washing (3x; S1), slices were transferred to a recording chamber and imaged with an upright microscope (Leica DMI6000FS, see above). We installed a custom-built reflective shield beneath the recording chamber for parallel monitoring of fluorescence and reflected light. At 1 Hz imaging cycles, we thus recorded two 510 nm fluorescence images (340 / 380 nm excitation) and a ‘pseudo-brightfield’ reflected light image that allowed quasi simultaneous analysis of intracellular Ca2+ and tubular movement.
Whole-mount seminiferous tubule imaging
Isolated tubules (>1 cm length) were placed onto a membrane within a custom-built 3D printed two-compartment recording chamber that was constantly superfused with S1. Small membrane holes under the tubules and around a defined stimulation area allowed for (i) gentle fixation of the tubules and (ii) focal ATP perfusion of selected tubular regions by vacuum-generated negative pressure (80-180 mmHg) in the submembraneous chamber compartment and continuous suction of S1 from the top compartment. After visual determination of tubular stage (I – III) (Parvinen, 1982), the perfusion pencil was positioned to selectively stimulate an area of known and homogeneous stage. Focal stimulation in the desired area was routinely confirmed by transient dye perfusion (Fast Green) prior to ATP exposure. ATP stimulations (100 μM; 10 s) and corresponding negative controls were compared to determine ATP-dependent Ca2+ signals (offspring from crossing SMMHC-CreERT2 and Ai95D mice) or tubular contractions and sperm transport. For low-magnification brightfield or fluorescence imaging, we used a MacroFluo Z16 APO A system (Leica Microsystems) equipped with either a DFC450C camera and a PLANAPO 1.0x / WD 97 mm objective (brightfield) or with a monochrome DFC365FX camera and a 5.0x/0.50 LWD PLANAPO objective (fluorescence). Images were acquired at 1 Hz.
In vivo imaging
We administered tamoxifen (75 mg tamoxifen kg−1 body weight) to double-positive adult male offspring (SMMHC-CreERT2 x Ai95D) via daily intraperitoneal injections for 5 consecutive days. Mice were closely monitored for any adverse reactions to the treatment. Experiments were performed 2–5 weeks after the first injection. For surgery, mice were anesthetized with ketamine-xylazine-buprenorphine (100 mg kg−1, 10 mg kg−1, 0.05–0.1 mg kg−1, respectively; Reckitt Benckiser Healthcare, UK). First, we made an incision next to the linea alba in the hypogastric region, followed by a 5 mm incision into the peritoneum. One testis was gently lifted from the abdominal cavity. Its gubernaculum was cut and the testis – with the spermatic cord, its blood vessels and vas deferens still intact – was transferred to a temperature-controlled imaging chamber filled with extracellular solution (S1; 35°C), mounted on custom-designed 3D printed in vivo stage (Figure 7–figure supplement 1). Throughout each experiment, vital signs (heartbeat, blood oxygen level, breathing rhythm) were constantly monitored and recorded (breathing). Moreover, we routinely checked unobstructed blood flow within testicular vessels during experiments. To avoid movement artifacts, the tunica was glued to two holding strings using Histoacryl tissue adhesive. After surgery, anesthesia was maintained by constant isoflurane inhalation (1–1.5% in air). Time-lapse intravital imaging was performed using a Leica TCS SP8 MP microscope. For incident light illumination / reflected light widefield recordings (5–10 Hz), we used N PLAN 5x/0.12 or HC APO L10x/0.30 W DLS objectives with large fields of view. Multiphoton time-lapse images were acquired at ~2 Hz frame rates using external hybrid detectors and the HCX IRAPO L25x/0.95 W objective at 930 nm excitation wavelength. For in vivo stimulation experiments, we used a Picospritzer III (Parker Hannifin, Pine Brook, NJ) to puff nanoliter volumes of control saline (S1; containing Alexa Fluor 555 (4 μM)) or stimulus solution (S1; containing Alexa Fluor 555 (4 μM) and ATP (1 mM)), respectively, from beveled glass micropipettes onto the surface of seminiferous tubules.
Data analysis
All data were obtained from independent experiments performed on at least three days. Individual numbers of cells / tubules / experiments (n) are denoted in the respective figures and / or legends. If not stated otherwise, results are presented as means ± SEM. Statistical analyses were performed using paired or unpaired t-tests, one-way ANOVA with Tukey’s HSD post hoc test or the Fisher Exact test (as dictated by data distribution and experimental design). Tests and corresponding p-values that report statistical significance (≤0.05) are individually specified in the legends. Data were analyzed offline using FitMaster 2.9 (HEKA Elektronik), IGOR Pro 8 (WaveMetrics), Excel 2016 (Microsoft, Seattle, WA), and Leica LAS X (Leica Microsystems) software. Dose-response curves were fitted by the Hill-equation. Time-lapse live-cell imaging data displaying both Ca2+ signals and tubular contractions were analyzed using custom-written code in MATLAB (The MathWorks, Natick, MA).
For quantitative image analysis, images from both reflected light and fluorescence time-lapse recordings were registered to their respective first image frame at time point t0, using the registration algorithm from (Liu et al., 2015) (implementation in (Evangelidis, 2013)), resulting in stabilized recordings without movement. For fura-2 fluorescence recordings, we first performed a single registration on the combined image (f340 + f380) and then applied the displacement vector field, computed by the registration algorithm, to both images (f340 and f380) separately. ROIs were defined manually at t0 and superimposed onto all subsequent images of the stabilized recording. At each time point ti, the fluorescence signal F was computed as the mean f340/f380 ratio of all pixels within a given ROI. When measuring Ca2+-dependent changes in GCaMP6f intensity, the fluorescence signal F was normalized with respect to a baseline before stimulation, computing the intensity change for the ith time point as . For clarity, linear baseline shifts were corrected in some example traces.
Seminiferous tubule contractions and transport of luminal content were visualized by reflected light microscopy of acute slices or whole-mount macroscopic tubule imaging, respectively. Data from both types of time-lapse recordings were analyzed and quantified as either flow strength or flow change (see below). For each frame at a given time point ti, the registration algorithm computed a flow or displacement vector field , where v1,1 = (x, y) is a vector indicating strength and direction of the displacement of pixel (1,1) between time points t0 and ti. The average norm is a measure for the effort that is necessary to register the image at t0 to the image at ti. The flow field strength quantified by this measure is interpreted as the amount of visible changes that, dependent on the experiment, result from tubule contraction and / or luminal content movement. For analysis of contractions in acute seminiferous tubule slices (Figures 3, 4, 7), we quantified the flow strength si within an ROI as the average norm |Vi| computed only for the vp,q corresponding to pixels within the ROI defined at t0. For whole-mount macroscopic imaging of luminal content movement in intact tubule segments (Figure 5), we quantified the flow change ci = si − si−1 as the change of flow strength between two consecutive time points / frames. Here, si values were preprocessed by smoothing with a moving average filter. Results are reported as the AUC, i.e., the area under the ci curve.
For analysis of in vivo data, we employed a custom set of ImageJ macros utilizing build-in functions of Fiji-ImageJ (Rueden et al., 2017; Schindelin et al., 2012). Widefield imaging data was first corrected for brightness fluctuation caused by a 50 Hz AC power supply. Here, we used the bleach correction plugin in histogram matching mode (Miura et al., 2014). Next, we applied Gaussian filter functions (GausBlur (5 px radius) and Gaussian Blur3D (x=0, y=0, z=5)). We calculated flow change via the Gaussian Window MSE function (sigma = 1; max distance = 3). Tubule selection used the polyline tool (line width adjusted to tubule diameter). Selected tubules ranged from 200 μm to 3.4 mm length. Next, flow fields of individual tubules were straightened. Average movement intensity was calculated from transversal line profiles (perpendicular to the straightened longitudinal axis of each tubule) and plotted as kymographs (space-time plots) to measure movement progression speed from linear regressions.
Multiphoton time-lapse imaging data was recorded in dual-channel mode, with (i) a target channel recording GCaMP6f fluorescence and some background signal (525\50 nm), and (ii) a background channel mainly recording autofluorescence (585\40 nm), allowing for background correction of the GCaMP6f signal using a dye separation routine. Slow constant movement in both channels was registered and removed to correct for steady drift. After Gaussian filtering (GausBlur (5 px radius); Gaussian Blur3D (x=0, y=0, z=5)), flow fields were calculated from the background signal. Again, flow change was calculated via the Gaussian Window MSE function (sigma = 1; max distance = 3).
Additional information
Competing interests
The authors declare no competing financial interests.
Funding
This work was funded by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) – 368482240/GRK2416 (NM & MSp); 412888997 (DF); 245169951 (AMa & MSp) – and by the Volkswagen Foundation (MSp, I/83533); MSp is a Lichtenberg Professor of the Volkswagen Foundation.
Author contributions
D.F. and L.K. contributed equally and have equal right to list themselves first in bibliographic documents. D.F., L.K., A.Ma., J.S., and M.Sp. contributed substantially to the conception of this work. D.F., L.K., N.M., N.U., J.S., and M.Sp. designed experiments. D.F., L.K., N.M., F.B., A.Mi., R.M., N.U., M.St., and J.S. contributed to data acquisition and analysis. D.F., L.K., N.M., M.St., F.B., A.Ma., J.S., and M.Sp. contributed to data interpretation. M.St. and D.M. created new software used for contraction analysis. All authors have drafted the work and / or substantively revised it. All authors have approved the submitted version of the manuscript.
Ethics
Mice were maintained and sacrificed according to European Union legislation (Directive 2010/63/EU) and recommendations by the Federation of European Laboratory Animal Science Associations (FELASA). All experimental procedures were approved by the State Agency for Nature, Environment and Consumer Protection (LANUV; protocol number / AZ 84-02.04.2016.A371).
Data and materials availability
All data is available in the main text or the supplementary materials. Previously unpublished source code for data analysis (quantification of tubular contractions, flow strength/change, Ca2+ signals) is available at: https://github.com/rwth-lfb/Fleck_Kenzler_et_al
Movie S1: SMMHC-CreERT2 mice allow inducible TPC-specific expression of genetically encoded fluorescent reporter proteins. After tamoxifen injections, SMMHC-CreERT2 x Ai14D male offspring express tdTomato (red) in both TPCs and vascular smooth muscle cells. Movie shows the 3D reconstruction of an intact and cleared (CLARITY (Chung and Deisseroth, 2013)) 6 × 3 × 1.5 mm testis sample with nuclei labelled by DRAQ5 (blue).
Movie S2: Quasi-simultaneous recording of peritubular Ca2+ signals and seminiferous tubule movement. A representative seminiferous tubule section (250 μm) is stimulated with ATP (100 μM, 10 s). After fura-2 bulk loading, ratiometric fluorescence imaging (f340/f380) reveals relative changes in Ca2+ concentration (rainbow colour map; blue, low Ca2+; red, high Ca2+) in a peripheral band of putative TPCs at the tubule’s edge. Since each image acquisition cycle (1 Hz) captures two fluorescence (Exλ340; Exλ380) and one reflective light image (brightfield), time-lapse recordings allow parallel physiological phenotyping of both seminiferous tubule Ca2+ responses and movement (shown sequentially for clarity).
Movie S3: Both ATP-induced seminiferous tubule Ca2+ responses and contractions are dose-dependent. A representative seminiferous tubule section (250 μm; fura-2 bulk loading) is stimulated with increasing ATP concentrations (1–1000 μM, 10 s). Ratiometric fluorescence imaging (f340/f380) reveals relative changes in Ca2+ concentration (rainbow colour map; blue, low Ca2+; red, high Ca2+) in putative TPCs. Quasi-simultaneous time-lapse recording of fluorescence (Exλ340; Exλ380) and brightfield (reflective light) images illustrates that both seminiferous tubule Ca2+ signals and contractions (shown sequentially for clarity) are dose-dependent and share an ATP threshold concentration of approximately 1 μM.
Movie S4: ATP stimulation triggers movement of luminal content in intact seminiferous tubules. Brightfield time-lapse recording of an intact isolated seminiferous tubule (field of view shows cycle stages II and III) challenged by brief focal ATP perfusion (100 μM, 10 s). The spatial extent of the stimulation zone had been defined by prior perfusion with a dye solution (food color).
Movie S5: ATP stimulation triggers transient Ca2+ signals in TPCs of intact seminiferous tubules. Fluorescence time-lapse recording of an intact seminiferous tubule (field of view shows cycle stage II) isolated from a mouse selectively expressing GCaMP6f in TPS (SMMHC-CreERT2 x Ai95D male offspring). Fluorescence imaging (ΔF/F) during brief focal ATP perfusion (100 μM, 10 s) – the spatial extent of the stimulation zone had been defined by prior perfusion with a dye solution (food color) – reveals relative changes in TPC Ca2+ concentration (rainbow colour map; blue, low Ca2+; red, high Ca2+).
Movie S6: In vivo multiphoton microscopy demonstrates spontaneous Ca2+ signals in mouse TPCs. Spontaneous seminiferous tubule in vivo activity monitored in SMMHC-CreERT2 x Ai95D mice. Intravital multiphoton fluorescence time-lapse imaging (ΔF/F, 2 Hz) reveals coordinated changes in TPC Ca2+ concentration (rainbow colour map; blue, low Ca2+; red, high Ca2+) among one of three seminiferous tubules in the field of view (591 μm × 591 μm).
Movie S7: Coordinated contractile movements ensure luminal sperm transport in vivo. Intravital en-face brightfield imaging illustrates spontaneous contractions and luminal movement in seminiferous tubules of adult mice. Low (1.5 mm × 1.4 mm field of view) and high-magnification time-lapse recordings reveal that contractions and luminal content propulsion are routinely observed in vivo. Note the unobstructed blood flow within testicular vessels.
Movie S8: Focal ATP stimulation triggers peritubular Ca2+ signals and seminiferous tubule contractions in vivo. Intravital multiphoton fluorescence time-lapse imaging in SMMHC-CreERT2 x Ai95D mice. Overlay of two detection channels (ΔF/F, GCaMP6f, green; Alexa Fluor 555, red). Stimulus solution (containing Alexa Fluor 555 (4 μM) and ATP (1 mM)) is puffed from a glass micropipette, which penetrated the tunica albuginea to target the interstitial space. Changes in TPC Ca2+ concentration are color-coded (black, low Ca2+; green, high Ca2+). Note that typically such contractions / Ca2+ signals do not occur when ATP is omitted from the ‘puff’ solution (data not shown).
Acknowledgements
We thank Corinna Engelhardt, Jessica von Bongartz, and Stefanie Kurth (RWTH Aachen University) for assistance, Andreas Meinhardt and Jörg Klug (Justus-Liebig-University Giessen) for kindly providing detailed information about the mouse TPC cell culture protocol, Andrea Mietens and Ralf Middendorff (Justus-Liebig-University Giessen) for comments and suggestions, J. Ullrich Schwarzer (Andrology-Center, Munich) and Frank-Michael Köhn (Andrologicum, Munich) for providing human samples, and all members of the Spehr laboratory for discussions.