ATP activation of peritubular cells drives testicular sperm transport

Spermatogenesis, the complex process of male germ cell proliferation, differentiation, and maturation, is the basis of male fertility. 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 toward the rete testis. 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.


Introduction
Spermatogenesis ranks among the most complex, yet least understood developmental processes in postnatal life. Inside the seminiferous tubules, which represent the functional units of the testis, this intricate course of mass cell proliferation and transformation events generates haploid spermatozoa from diploid spermatogonial stem cells. The seminiferous epithelium is composed of Sertoli cells, each intimately associated with !30 germ cells at different developmental stages (Mruk and Cheng, 2004). Sertoli cells provide the microenvironment critical for spermatogenesis by establishing the blood-testis barrier (Cheng and Mruk, 2010), forming the spermatogonial stem cell niche (Oatley and Brinster, 2012), and controlling epithelial cyclicity via auto-, para-, and endocrine feedback (Heindel and Treinen, 1989). Different types of spermatogonia (type A, intermediate, type B) are localized along the seminiferous tubule basement membrane (Chiarini-Garcia and Russell, 2001). Upon detachment, type B spermatogonia enter meiosis as preleptotene spermatocytes. During meiotic divisions and subsequent maturation steps, germ cells progress from primary to secondary spermatocytes and round to elongated spermatids.
Along the seminiferous epithelium, spermatogenesis has been conceptualized by attribution of sequential cellular 'stages' (Figure 1A), which progress through coordinated cycles (Hess and De Franca, 2008;Russell, 1990). First initiated in mice about 7 days postpartum (Kolasa et al., 2012), each spermatogenic cycle comprises 12 stages and lasts 8.7 days (Hermo et al., 2010). After approximately 39 days (4.5 cycle repetitions), spermatogenesis completes with the release of immotile spermatozoa from the seminiferous epithelium into the lumen of the tubule (spermiation). 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. eLife digest As sperm develop in the testis, the immature cells must make their way through a maze of small tubes known as seminiferous tubules. However, at this stage, the cells do not yet move the long tails that normally allow them to 'swim'; it is therefore unclear how they are able to move through the tubules. Now, Fleck, Kenzler et al. have showed that, in mice, muscle-like cells within the walls of seminiferous tubules can create waves of contractions that push sperm along. Further experiments were then conducted on cells grown in the laboratory. This revealed that a signaling molecule called ATP orchestrates the moving process by activating a cascade of molecular events that result in contractions. Fleck, Kenzler et al. then harnessed an advanced microscopy technique to demonstrate that this mechanism occurs in living mice. Together, these results provide a better understanding of how sperm mature, which could potentially be relevant for both male infertility and birth control.  (Hess and De Franca, 2008;Russell, 1990), which are arranged in consecutive order along the length of the tubule. A single layer of flat testicular peritubular cells (TPC; green) lines the tubular wall. Sertoli cells (SC) span the tubule from the basal lamina to the lumen. Developing germ cells (GC) are distributed between Sertoli cells. Spermatogonia are located near the basal membrane. Prophase spermatocytes move across the bloodtestis barrier (BTB) to the adluminal compartment where they complete meiosis. The resulting haploid spherical cells (round spermatids) differentiate into elongated spermatids and, eventually, into highly condensed and compartmentalized spermatozoa (spermiogenesis). These mature, yet still immotile germ cells are then released into the lumen (spermiation). E, epididymis; TA, tunica albuginea; inspired by Hess and De Franca, 2008. (B) Immunostaining against a-smooth muscle actin (a-SMA, green) marks TPCs in vitro (Tung and Fritz, 1989). Cell count is determined by nuclear staining (DAPI, blue). Cultures retain high TPC purity for at least two passages (92 ± 2%, n = 1102 (#1); 86 ± 3%, n = 542 (#2)). Dashed  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 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 mouse seminiferous tubules both in vitro and in vivo.

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., 2011;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 1B, and Figure 1-figure supplement 1A&B) and cells express transcripts for several ionotropic (P2X2, P2X4, P2X7) and metabotropic (P2Y2, P2Y6) purinoceptors ( Figure 1C). The specific biophysical and pharmacological profile of ATP-dependent transmembrane currents ( Figure 1D-H) strongly suggests functional expression of P2X2 and/or P2X4, but not P2X7 receptors. As reported for both P2X2 and P2X4 (North, 2002), TPC currents are saturated at 100 mM ATP ( Figure 1D&E), Figure 1 continued depicting a TPC (dashed line) targeted by a patch pipette (pp). (D II -D IV ) Original whole-cell recordings illustrate representative currents in response to ATP stimulation (100 mM (D II ) vs. 1000 mM (D III ) and 10 s (D III ) vs. 30 s (D IV ), respectively). V hold = À80 mV. (E, F) Quantification (bar charts; mean ± SEM; n as indicated) reveals saturation of peak current density at 100 mM ATP (E) and modest desensitization at a concentration-dependent rate (F). (G) Whole-cell voltage-clamp recordings show ATP-induced currents (100 mM; 10 s) that are potentiated by ivermectin (3 mM) and partially inhibited by suramin (100 mM), respectively (!60 s preincubation). V hold = À80 mV. (H) Quantification (bar charts; mean ± SEM; data normalized to initial control response) demonstrates dichotomy in drug sensitivity. Treatment was categorized as effective (eff) if current amplitudes deviate by ± SD from average control recordings (85 ± 24%, 2 nd ATP stimulation). Note that each drug proved ineffective (no eff) in some cells. Gray circles depict data from individual cells. (I-L) ATP-dependent Ca 2+ mobilization in cultured TPCs. Ca 2+ transients in response to repetitive stimulation (10 mM, 10 s) are monitored by ratiometric (fura-2) fluorescence imaging. (I) Phase contrast (I) and merged fluorescence (f 380 ; II) images of a TPC in vitro. Bottom pseudocolor frames (rainbow 256 color map) illustrate relative cytosolic Ca 2+ concentration ([Ca 2+ ] c ) before (III) and during (IV) ATP stimulation. (J, K) Representative original traces from time-lapse fluorescence ratio (f 340 /f 380 ) recordings depict repetitive [Ca 2+ ] c elevations upon ATP exposure under control conditions ((J) blue traces correspond to the TPC in (I)) and during reduced extracellular Ca 2+ concentration ((K) [Ca 2+ ] e = 100 nM; 60 s preincubation). (L) Bar chart depicting Ca 2+ signal amplitudes (mean ± SEM; n as indicated) -normalized to the initial ATP response -under control conditions (gray) vs. low [Ca 2+ ] e (green). Asterisks denote statistically significant differences (* 1 p=0.001; * 2 p=0.002; * 3 p=5.5e À5 ; * 4 p=0.0006; * 5 p=0.02; * 6 p=0.02; Student t-test (F, L), one-way ANOVA (H)). (M) Representative whole-cell voltage-clamp recordings (V hold = À80 mV) of ATP-induced inward currents in cultured mouse TPCs. Two components -a fast relatively small current (arrow head) and a delayed lasting current (asterisk) -are triggered repeatedly by successive ATP exposure (100 mM; 90 s inter-stimulus interval). Notably, we never observed a delayed slow current without a fast response. (N) Bar chart quantifying peak densities (mean ± SEM, circles show individual values) of the fast (I fast ; black) and the delayed (I slow ; gray) ATP-induced current components (1 st stimulation: I fast 0.37 ± 0.2 pA/pF; I slow 1.58 ± 0.5 pA/pF; 2 nd stimulation I fast 0.27 ± 0.1 pA/pF; I slow 1.37 ± 0.4 pA/pF). (O) Bar graph illustrating the frequency of I fast (black) and I slow (gray) occurrence upon ATP (100 mM) stimulation in absence (w/o) and presence (w) of GTP (500 mM) in the pipette solution, respectively. Asterisks denote statistically significant differences (* 7 p=0.008, * 8 p=0.0003; Fisher's exact test); n as indicated in bars. (P) Representative whole-cell voltage-clamp recordings (V hold = À80 mV) of inward currents induced by ATP (100 mM) and UTP (100 mM), respectively. Whenever ATP triggers both I fast and I slow (left), I slow is also induced by UTP (right). UTP-dependent currents develop significantly slower than ATPevoked I fast (inset; * 9 p=0.03; paired t-test). The online version of this article includes the following figure supplement(s) for figure 1:   . ATP triggers TPC Ca 2+ signals in acute seminiferous tubule sections. (A) 3D reconstruction of an intact 6 Â 3 Â 1.5 mm testis sample from a SMMHC-CreER T2 x Ai14D mouse after tissue clearing (CLARITY Chung and Deisseroth, 2013) reveals tdTomato expression (red) restricted to TPCs and vascular endothelial cells (asterisk in (A II )). Nuclear staining (DRAQ5; blue) is most prominent in post-meiotic germ cells because of their high degree of DNA condensation. (B) SMMHC-CreER T2 -expressing cells in the tubule wall are TPCs. In testis cryosections from adult SMMHC-CreER T2 x Ai14D mice, Cre-driven tdTomato signals (B I ) and a-SMA immunostaining (B II ) colocalize at tubular margins (B III ). Nuclei are stained with DRAQ5 (blue (B IV )). Note that endothelial vasculature in interstitial regions (asterisks) is a-SMA-negative (merged image (B V )). (C-H) Both TPC-specific expression of a genetically encoded Ca 2+ indicator (GCaMP6f) and bulk loading with a synthetic Ca 2+ sensor (fura-2) allow for TPC-selective live cell Ca 2+ imaging in acute seminiferous tubule sections. (C) Merged fluorescence and reflected light micrographs show the location of SMMHC-expressing TPCs (red) within the wall of an intact tubule. (D-E) Cre-dependent GCaMP6f expression in SMMHC-CreER T2 x Ai95D mice reveals Ca 2+ transients in TPCs in response to ATP. Representative fluorescence images ((D) rainbow 256 color map) before and during ATP exposure (100 mM; 10 s), and corresponding traces (E) showing changes in GCaMP6f intensity (DF/F) over time. Traces from ROIs outlined in (D) (white solid lines). (F) Merged fluorescence image of an acute seminiferous tubule section from an AMH-Cre x Ai14D mouse after bulk loading with fura-2/AM (green). Anti-Mü llerian hormone (AMH) dependent expression of tdTomato (red) specifically labels Sertoli cells that build the seminiferous epithelium. Note the narrow green band of marginal TPCs that are preferentially labeled by the Ca 2+ -sensitive dye. (G-H) Ratiometric Ca 2+ imaging in fura-2-loaded tubules enables semi-quantitative live-cell Figure 2 continued on next page whereas P2X7 receptors display strongly reduced ATP sensitivity (Donnelly-Roberts et al., 2009). Moreover, currents recorded from TPCs showed modest but persistent desensitization ( Figure 1D&F), which is similarly observed for recombinant P2X2 and P2X4, but not P2X7 receptors (Coddou et al., 2011). TPCs also displayed reduced BzATP sensitivity (data not shown), which is a potent activator of P2X7 receptors (Donnelly-Roberts et al., 2009). Ivermectin ( Figure 1G&H), an agent selectively potentiating P2X4 receptor currents (Khakh et al., 1999;Silberberg et al., 2007), increased ATP-induced currents in a subpopulation of TPCs (n = 7/12), whereas suramin ( Figure 1G&H), a drug inhibiting P2X2, but not P2X4 receptors (Evans et al., 1995), inhibited a TPC subset (n = 10/18).
Notably, live-cell ratiometric Ca 2+ imaging in cultured TPCs revealed robust and repetitive cytosolic Ca 2+ transients upon ATP exposure ( Figure 1I&J). We next reduced the extracellular Ca 2+ concentration ([Ca 2+ ] e ) to 100 nM, a concentration approximately equimolar to cytosolic levels, by adding an appropriate chelator/ion concentration ratio (1 mM EGTA/0.5 mM CaCl 2 ). This treatment, which drastically diminishes the driving force for Ca 2+ influx, did substantially reduce, but not abolish ATP response amplitudes ( Figure 1K&L). The selective P2Y receptor agonist UTP (Alexander et al., 2019b) also triggered Ca 2+ signals (data not shown), indicating a role for G protein-dependent Ca 2+ release from internal storage organelles (Müller et al., 2020). 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 1M&N). We hypothesized that this slower current could result from P2Y receptor-/G protein-dependent Ca 2+ release, likely mediated by the P2Y2 isoform since P2Y6 receptors lack substantial ATP sensitivity (Alexander et al., 2019a;Jacobson et al., 2015). Indeed, occurrence of the delayed current depends on presence of intracellular GTP ( Figure 1O). Moreover, selective recruitment of G protein-coupled P2Y receptors with UTP ( Figure 1P) exclusively triggered such slowly developing currents. Largely carried by Cl - (Figure 1-figure supplement 2), this current likely results from P2Y receptor-mediated phosphoinositide turnover, Ca 2+ release, and activation of Ca 2+ -gated Clchannels. 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. Therefore, we examined purinergic Ca 2+ signals from mouse TPCs in acute seminiferous tubule sections (Fleck et al., 2016). In parallel approaches, we employed two different fluorescent Ca 2+ reporters, a synthetic ratiometric Ca 2+ sensor (fura-2) as well as a genetically encoded Ca 2+ indicator (GCaMP6f). The dual excitation ratiometric indicator fura-2 allows semi-quantitative Ca 2+ measurements (Bootman et al., 2013), but lacks cell type specificity as tubules are bulk-loaded with a membrane-permeable acetoxymethyl ester conjugate. By contrast, conditional gene targeting via the Cre/Lox system (Smith, 2011) allows TPCspecific expression of the single-wavelength indicator GCaMP6f. First, we confirmed inducible TPC-targeted testicular expression of fluorescent reporter proteins in SMMHC-CreER T2 x Ai14D mice ( Figure 2A-C, Video 1). Tamoxifen-induced transgenic expression of CreER T2 under control of the mouse smooth muscle myosin, heavy polypeptide 11 (a.k.a. SMMHC) promoter drives Cremediated recombination of loxP-flanked reporters (tdTomato (Ai14D mice) or GCaMP6f (Ai95D)) Corresponding traces (H) show the fluorescence intensity ratio (f 340 /f 380 ) from four ROIs (in (G); white solid lines) over time. Inset (G) shows a putative TPC and an adjacent putative spermatogonium (asterisk) at higher magnification.
Video 1. SMMHC-CreER T2 mice allow inducible TPCspecific expression of genetically encoded fluorescent reporter proteins. After tamoxifen injections, SMMHC-CreER T2 x Ai14D male offspring express tdTomato (red) in both TPCs and vascular smooth muscle cells. Video shows the 3D reconstruction of an intact and cleared (CLARITY, Chung and Deisseroth, 2013)  in smooth muscle cells and TPCs (Wirth et al., 2008). Second, TPC-specific GCaMP6f expression in SMMHC-CreER T2 x Ai95D mice revealed robust Ca 2+ transients in cells of the tubular wall upon ATP exposure ( Figure 2D&E). Third, fura-2/AM loading of acute seminiferous tubule sections preferentially labeled the outermost cell layer ( Figure 2F), allowing semi-quantitative in situ imaging of ATPdependent Ca 2+ signals in mouse TPCs ( Figure 2G&H). So far, our results thus demonstrate that challenging TPCs with extracellular ATP triggers robust Ca 2+ signals both in vitro and in situ.

ATP triggers seminiferous tubule contractions
We hypothesized that ATP-induced Ca 2+ 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 Ca 2+ responses and seminiferous tubule movement (methods). Brief ATP exposure resulted in a peripheral band of Ca 2+ activity at the edge of the tubule. Such signals usually coincided with a pronounced contractile motion of the seminiferous tubule ( Figure 3A, Video 2). When movement is quantified as the time-lapse image flow field  strength (methods) tubular contraction follows the Ca 2+ signal onset with minimal delay, outlasts the Ca 2+ signal peak, and recovers slowly ( Figure 3B). Both Ca 2+ responses and tubular movement are dose-dependent and share an ATP threshold concentration of approximately 1 mM ( Figure 3C-E, Video 3). Contractile smooth muscle plasticity (Tuna et al., 2012) likely underlies the apparent difference in signal saturation ( Figure 3D). Notably, in some tubules, we observed spontaneous low-Video 2. Quasi-simultaneous recording of peritubular Ca 2+ signals and seminiferous tubule movement. A representative seminiferous tubule section (250 mm) is stimulated with ATP (100 mM, 10 s). After fura-2 bulk loading, ratiometric fluorescence imaging (f 340 /f 380 ) reveals relative changes in Ca 2+ concentration (rainbow color map; blue, low Ca 2+ ; red, high Ca 2+ ) in a peripheral band of putative TPCs at the tubule's edge. Since each image acquisition cycle (1 Hz) captures two fluorescence (Exl 340 ; Exl 380 ) and one reflective light image (brightfield), time-lapse recordings allow parallel physiological phenotyping of both seminiferous tubule Ca 2+ responses and movement (shown sequentially for clarity).

Figure 4 continued
tubule contractions with only minor response adaptation. Traces depict changes in fura-2 intensity ratio (f 340 /f 380 ; black) or tubular movement (red) upon brief ATP exposure (10 s; 10 mM; 5 min intervals) under control conditions. (C) Reducing [Ca 2+ ] e (100 nM; 5 min incubation) strongly diminishes responses to ATP (10 s; 10 mM). (D) Depletion of internal Ca 2+ stores (CPA Seidler et al., 1989; 90 mM; 18.8 ± 9.3 min incubation) essentially abolishes both Ca 2+ signals and tubule contractions. (E-G) Quantification of data exemplified in (B-D). (E) Bar chart depicting ATP sensitivity (response rate; %), independent of signal strength. Occurrence of Ca 2+ elevations (black) and tubule contractions (red) are plotted for different experimental conditions [i.e. stimulation with ATP or UTP, regular or reduced [Ca 2+ ] e (1 mM or 100 nM, respectively), and Ca 2+ indicator (fura-2 or GCaMP6f, respectively)]. Numbers of experiments are indicated in each bar. (F) Signal amplitudes (Ca 2+ , black; contractions, red) of responding TPCs/tubules, quantified as a function of stimulus, treatment, and Ca 2+ sensor. Data (mean ± SEM) are normalized to the respective initial responses to ATP (10 mM) under control conditions (dotted horizontal line; see first stimulations in B and C). Experimental conditions and numbers of experiments as in (E). Asterisks denote statistical significance (* 1 p=2.2e À19 and * 2 p=5.6e À8 ; t-test; note: tests only performed when n > 5 and only one variable was changed). (G) Pie charts illustrating the profoundly reduced ATP sensitivity of TPCs/tubules after depletion of Ca 2+ storage organelles (CPA; 90 mM). Numbers within pies correspond the total count of cells/tubules that responded to ATP before treatment. (H) Effects of lowering [Ca 2+ ] e are comparable over both incubation periods and concentrations in the nanomolar range. Significantly reduced, though not abolished TPC Ca 2+ signals (left) and seminiferous tubule contractions (right) are observed in presence of both 100 nM and 12 nM [Ca 2+ ] e as well as for variable incubation periods lasting between 1 and 10 min, respectively. Asterisks denote statistical significance (* 3 p=2.8e À10 , * 4 p=7.8e À14 , * 5 p=3.8e À9 , * 6 p=1.0e À6 ; one-way ANOVA with post-hoc Tukey HSD test; note: tests only performed when n ! 5). The online version of this article includes the following figure supplement(s) for figure 4: 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 Ca 2+ signaling mechanism(s) underlying ATP-dependent TPC contractions. First, we asked whether influx of external Ca 2+ is involved in TPC force generation. Similar to in vitro observations ( Figure 1K&L), diminishing or even reversing the driving force for transmembrane Ca 2+ flux by reducing [Ca 2+ ] e to 100 nM or 12 nM, respectively, for variable durations, significantly decreased both TPC Ca 2+ signals and tubular contractions ( Figure 4B-H). While, upon [Ca 2+ ] e reduction, ATP-dependent responses (both Ca 2+ signals and contractions) were still detected in the vast majority of cells/experiments ( Figure 4E), response strength was strongly diminished ( Figure 4F). These effects were independent of both the extent (12 nM or 100 nM) and the duration (1-10 min) of [Ca 2+ ] e reduction and were fully reversible ( Figure 4H). Second, we examined a potential role of ATP-induced Ca 2+ release from internal storage organelles. Ca 2+ depletion of the Intact seminiferous tubules are placed on transparent foil in a custom-built macroscopic imaging chamber. The tubule is kept stationary by gentle suction through tiny holes punched in the foil and vacuum underneath. As previously suggested (Hess and De Franca, 2008), tubules are coarsely categorized into three stages (I-III; color code) according to luminal sperm content. Precise mapping of stimulated regions is feasible by positioning both tubule and perfusion pencil within an area delimited by several holes that outline a stimulation zone (methods). The tubule region directly exposed to ATP is designated as ROI 0, with adjacent equidistant sections numbered consecutively (up to ROI ±6; schematic). In the original image shown, only ROIs ± 3 are outlined for clarity. (B) Analysis of luminal content movement by calculation of flow change c i relative to each previous image (methods) within a representative luminal ROI. Motion is quantified by measuring the area under curve (AUC; solid red) within 60 s after stimulation onset. Note that mechanical control stimulation (extracellular solution) does not affect basal luminal motion. (C) Bar charts depicting luminal content movement (means ± SEM) upon ATP stimulation (100 mM; 10 s) in either directly exposed regions (C I ; n = 17) or adjacent areas (C II ; n = 3-17). Green/blue (groups I and III) and red (group II) bars depict stages with a low vs. a high luminal sperm count, respectively. Horizontal gray lines mark the average basal luminal motion prior to stimulation. ATP induces significantly increased content movement in directly stimulated areas (ROI 0) independent of luminal sperm count/stage group (C I ). Note that in adjacent regions (C II ) unidirectional movement occurs exclusively in tubule sections with high luminal sperm density. Asterisks denote statistically significant differences (* 1 p=8.7e À5 ; * 2 p=6.7e À7 ; * 3 p=0.005; * 4 p=0.002; * 5 p=0.03; unpaired two-tailed t-test).
sarcoplasmic reticulum via pharmacological inhibition of the sarco/endoplasmic reticulum Ca 2+ -ATPase by cyclopiazonic-acid (CPA) essentially abolished both ATP-dependent Ca 2+ signals and contractions ( Figure 4D), with very few cells/ tubules retaining some residual ATP sensitivity during CPA treatment ( Figure 4G). Importantly, all results from ratiometric fura-2 imaging were qualitatively indistinguishable from those obtained with genetically targeted GCaMP6f ( Figure 4E-G), showing that both approaches to TPC Ca 2+ measurement provide comparable results. Third, given the pronounced effect of pharmacological store depletion, we aimed to quantify the specific contribution of metabotropic purinoceptors to the overall ATP-mediated effect. The P2Y receptor-selective agonist UTP (Coddou et al., 2011) evoked both TPC Ca 2+ signals and tubular contractions ( Figure 4E&F). However, under control [Ca 2+ ] e conditions, UTPevoked responses were substantially reduced compared to control ATP stimulations ( Figure 4F). Notably, these UTP responses were statistically indistinguishable from the diminished ATP-dependent signals we observed under low [Ca 2+ ] e conditions ( Figure 4F).
Together, these data strongly suggest that (i) extracellular ATP acts as a potent TPC stimulus that triggers seminiferous tubule contractions in situ, that (ii) P2X and P2Y receptors act in concert to mediate TPC responses to ATP exposure, that (iii), while P2X receptor-dependent external Ca 2+ influx apparently boosts responses to ATP, P2Y receptor-mediated Ca 2+ mobilization from the sarcoplasmic reticulum is necessary to evoke TPC responses, and consequently -since store depletion essentially abolishes ATP-dependent signals -that (iv) influx of external Ca 2+ 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 (

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 spermatogenic cycle into three distinct stage groups following published protocols (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 that outlasted the presence of ATP for several tens of seconds ( Figure 5B, Video 4). Second, we analyzed if luminal movement depends on the tubule's cycle stage and, consequently, luminal sperm count. When we analyzed ATP-induced movement in directly stimulated luminal regions (each designated as region-of-interest (ROI) 0) and compared stage groups with a high (group II) vs. a relatively low (groups I and III) amount of luminal sperm, we observed no difference in stimulation-dependent motion (Figure 5C I ). 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 fluid flow propagates beyond the directly stimulated tubule section. When we analyzed luminal motion in equidistant tubule sections adjacent to the directly stimulated area ROI 0 ( Figure 5A), we found a significant, though relatively small bidirectional wave of propagating movement in stage groups I and III, which exhibit a low luminal sperm count (Figure 5C II ). Strikingly, we observed strong unidirectional luminal movement upon ATP stimulation of stage group II tubule sections which show high luminal sperm density associated with spermiation ( Figure 5C II ). In this stage group, luminal content is predominantly propelled toward areas of ascending spermatogenic cycle stages. These findings demonstrate directionality of sperm transport upon focal purinergic TPC stimulation in isolated seminiferous tubules.
As expected, ATP-induced tubule contractions also manifest as Ca 2+ signals in TPCs ( Figure 6A&B, Video 5). However, these Ca 2+ elevations appear to be limited to those areas directly exposed to ATP (ROI 0). We observed no such signals in adjacent tubule sections independent of the stimulated stage group or an ascending or descending stage direction ( Figure 6C&D). This finding indicates that, in the isolated seminiferous tubule, ATP acts as a local 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. However, local contractions generate sufficient force to move luminal content beyond the directly stimulated area and, in turn, directionality of flow along short-tomedium distances ( 600 mm; Figure 5C II ) is not critically dependent on peristaltic contractility.

ATP induces tubular contractions in vivo
To ultimately attribute a physiological role to ATP-dependent Ca 2+ 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 epifluorescence and multiphoton microscopy of the mouse testis.
Initially, we monitored spontaneous seminiferous tubule activity in SMMHC-CreER T2 x Ai95D mice. Multiphoton time-lapse imaging revealed spontaneous TPC Ca 2+ signals that typically accompanied strong tubule contractions ( Figure 7A&B, Video 6). Several characteristics emerged from quantitative analysis of these observations. First, during sufficiently long recording periods ( 30 min 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). These movements coincide with 'macroscopic' Ca 2+ waves that travel at comparable speed and direction (Figure 7-figure supplement 2D). Notably, the observed coordinated contractile movements provide sufficient force to ensure luminal sperm transport (Video 7).
Finally, we examined if brief focal ATP stimulation also triggers peritubular Ca 2+ signals and seminiferous tubule contractions in vivo. Therefore, we filled low resistance patch pipettes with fluorescently labeled ATP solution, penetrated the tunica albuginea, and targeted the interstitial space close to neighboring tubules ( Figure 7E, Video 8). Nanoliter puffs of ATP-containing test solution induced both Ca 2+ 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 2E). Taken together, in vivo recordings demonstrate that robust recurrent seminiferous tubule contractions (i) occur spontaneously, (ii) are driven by cytosolic Ca 2+ elevations in TPCs that propagate in a wave-like fashion, 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 Video 5. ATP stimulation triggers transient Ca 2+ 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 TPCs (SMMHC-CreER T2 x Ai95D male offspring). Fluorescence imaging (DF/F) during brief focal ATP perfusion (100 mM, 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 Ca 2+ concentration (rainbow color map; blue, low Ca 2+ ; red, high Ca 2+ ).
https://elifesciences.org/articles/62885#video5  . Red triangles mark passive movements, which occur upon contractions of adjacent tubules. Note the lack of a corresponding Ca 2+ signal. (C) Analysis of spontaneous tubular motion in vivo. Low magnification incident light image of the mouse testis (C I ) shows several superficial seminiferous tubule segments, testicular blood vessels (white asterisk; note that unobstructed blood supply (i.e. visualizing erythrocyte flow) is checked routinely), and a specific segment outlined by red dotted lines. After time-lapse imaging, this segment is digitally straightened (C II ) and subjected to motion analysis.   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 both 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.
The site(s)/cellular origin of testicular ATP release as well as the mechanism(s) that trigger ATP secretion in vivo currently remain elusive. The apparent absence of efferent nerve endings in the seminiferous tubules and interstitial tissue (Tripiciano et al., 1996) suggests that tubule contractility is under endo-/paracrine control. By contrast, autonomic innervation of the testicular capsule mediates smooth muscle cell contraction of the tunica albuginea, using ATP as a (co)transmitter (Banks et al., 2006). Regulated ATP release has been reported for both Sertoli and germ cells (Gelain et al., 2005;Gelain et al., 2003). Moreover, TPCs express P2Y6 receptors ( Figure 1C), which were reported to mediate ATP release upon activation (Carneiro et al., 2014). Thus, TPCs could themselves participate in regenerative nucleotide release. During spermatogenesis, apoptosis is a vital process (Print and Loveland, 2000). In fact, up to 75% of germ cells undergo apoptosis under physiological conditions (Huckins, 1978). This substantial germ cell loss is called 'density-dependent regulation' (Hess and De Franca, 2008). Since ATP release from apoptotic cells is well documented (Elliott et al., 2009) it is likely that cell densitydependent waves of apoptosis could regularly generate local ATP surges. We have previously shown that one result of seminiferous extracellular ATP elevation is signal amplification by increased ATP release (Fleck et al., 2016), although the mechanistic basis of this positive feedback pathway is yet unknown. Given (i) the robust cytosolic Ca 2+ transients observed in response to ATP exposure in various testicular cell types (Fleck et al., 2016;Gelain et al., 2005;Liévano et al., 1996;Veitinger et al., 2011;Walenta et al., 2018) and (ii) the usually millimolar ATP content in secretory vesicles (Bodin and Burnstock, 2001;Zhang et al., 2007), the most parsimonious explanation for ATP-induced ATP release would be an inevitable ATP 'co-secretion' upon any Ca 2+ -dependent exocytosis event. In addition, ATP release has been observed in several cell types as a result of mechanical deformation, shear stress, stretch, or osmotic swelling (Button et al., 2013) adding another putative mechanism of regenerative signaling in purinergic contraction control.
Notably, extracellular ATP is rapidly degraded by ecto-nucleotidases (Zimmermann et al., 2012), rendering its interstitial half-life relatively short and, thus, narrowing its paracrine radius to a few hundred micrometers (Fitz, 2007). Combined with its fast diffusion -approximately 1 mm in less than 10 ms (Khakh, 2001) -extracellular ATP bears all characteristics of a fast paracrine agent in testicular communication (Praetorius and Leipziger, 2009).
Excitation-contraction coupling in TPCs is poorly described. Our results strongly suggest that a combination of P2X (isoforms 2 and/or 4) receptor-dependent Ca 2+ influx and P2Y (likely isoform 2) receptor-activated phospholipase Cb-dependent Ca 2+ release from the sarcoplasmic reticulum -the latter being critical and resembling the recently reported mechanism of vascular smooth muscle cell contraction in small pulmonary veins (Henriquez et al., 2018) -provides the [Ca 2+ ] c elevation required for force generation (Berridge, 2008). P2X and P2Y receptors act on different time scales and display different ligand sensitivity, with EC 50 values in the nanomolar (P2Y) vs. micromolar (P2X) range (North, 2002). It is possible that the co-activation of an ionotropic (P2X) and a metabotropic (P2Y) signaling pathway serves functions analogous to the concomitant exposure to both ATP and noradrenaline in mesenteric artery smooth muscle. Here, activation of P2X1 receptors generates a small initial contraction that is followed by larger noradrenaline-induced contraction (Lamont et al., 2006;Lamont et al., 2003). Regarding TPC [Ca 2+ ] c elevation, our data suggest that P2X receptor activation also targets Ca 2+ release from internal stores, as their depletion inhibits excitation-contraction coupling entirely. Therefore, it is likely that P2X receptors act as signal boosters that mediate Ca 2+ -induced Ca 2+ release, possibly via activation of ryanodine receptors (Berridge, 2008). This way, the combined action of P2X and P2Y receptors might equip TPCs with a broader 'two-step' stimulus integration range.
Whole-mount imaging of isolated seminiferous tubules reveals propagation of luminal content that extends beyond the confines of the stimulated/contracted area and displays stage-dependent directionality. While peristaltic contractions are driven by propagating wave-like Ca 2+ signals in vivo, focal ATP stimulation appears insufficient to trigger a regenerative Ca 2+ wave in isolated tubules. We, thus, conclude that the observed directionality results from other, likely structural characteristics, for example anatomical features of stage group II and III tubules that favor a specific flow direction (increased tubule diameter and reduced luminal resistance along the stage II-to-III transition zone). We cannot, however, rule out that the use of large field-of-view/low numerical aperture objectives for 'macroscopic' imaging simply prevents the detection of low-amplitude Ca 2+ signal spread.
Translation of our findings from the mouse model to humans awaits further in-depth investigation. We have recently reported that ATP activates Ca 2+ signals in human TPCs in vitro (Walenta et al., 2018). Moreover, our present findings reveal ATP-induced contractions in cultured human TPCs. There are, however, notable differences between the human and the mouse tubular walls. While a single layer of TPCs surrounds the mouse seminiferous tubules, the human tubular wall architecture is more complex, containing several TPC layers, substantial amounts of extracellular matrix proteins, and immune cells (Mayerhofer, 2013). Impaired spermatogenesis in sub-/infertile men typically coincides with tubular wall remodeling and a partial loss of TPC contractility proteins has been reported in infertile men (Welter et al., 2013). Accordingly, interference with TPC contractility had been proposed as a promising strategy for human male contraception (Romano et al., 2005). However, a causal relationship between contractility (or the lack thereof) and male (in)fertility has never been established. In fact, seminiferous tubule contractions had, so far, never been observed in vivo and most in vitro reports were based on indirect and non-quantitative evidence, for example from posthoc fluorescence or scanning electron microscopy (Barone et al., 2002;Fernández et al., 2008;Losinno et al., 2016;Losinno et al., 2012;Tripiciano et al., 1999;Tripiciano et al., 1997;Tripiciano et al., 1996), morphometry of single cells in culture (Rossi et al., 2002;Santiemma et al., 2001;Santiemma et al., 1996;Tripiciano et al., 1996), or intraluminal pressure analysis (Miyake et al., 1986;Yamamoto et al., 1989). The fact that expression of TPC contractility proteins initiates with puberty under androgen control and that selective androgen receptor knockout in TPCs renders mice infertile (Welsh et al., 2009) underscores a potential role of TPC contractions in male fertility. Accordingly, pharmacological targeting of purinergic signaling pathways to (re) gain control of TPC contractility represents an attractive approach for male infertility treatment or contraceptive development.
Among several remaining questions, future experimental efforts will have to address (i) whether TPCs are coupled by gap junctions to display coordinated activity; (ii) whether and, if so, how the final ATP metabolite adenosine affects seminiferous tubule physiology; (iii) whether Rho/Rho kinase signaling pathways modulate TPC contractility as frequently observed in other smooth muscle cells (Somlyo and Somlyo, 2003); (iv) what, if any, role is played by P2X receptor-dependent changes in membrane potential; (v) which function is served by the sustained Ca 2+ -gated Clcurrent (Figure 1figure supplement 2); (vi) why periods of enhanced contractile activity are interrupted by longer quiescent episodes (Figure 7-figure supplement 2B); (vii) which additional or complementary roles in TPC physiology are played by previously proposed activators, including vasopressin, oxytocin, prostaglandins, and endothelin (Albrecht et al., 2006;Mayerhofer, 2013); (viii) whether an additional cytosolic and/or membrane Ca 2+ oscillator (Berridge, 2008) provides an endogenous pacemaker mechanism that acts independent of purinergic stimulation; and (ix) whether, similar to vascular smooth muscle cells, some specific tone is maintained between contractions by spatial averaging of asynchronous oscillations (Berridge, 2008), a mechanism that could explain the occurrence of spontaneous low-amplitude 'vibratory' movements and local indentations that we ( Figure 4A) and others (Ellis et al., 1981;Worley et al., 1985) have observed.

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 hr light-dark cycle; food and water available ad libitum).

Stimulation
For focal stimulation, solutions and agents were applied from air pressure-driven reservoirs via an 8in-1 multi-barrel 'perfusion pencil' (AutoMate Scientific; Berkeley, CA). Changes in focal superfusion (Veitinger et al., 2011) 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 (Picospritzer III; Parker Hannifin, Hollis, NH).
Low [Ca 2+ ] e solutions (S 3 and S 4 ) were applied via both the bath and perfusion pencil. To ensure depletion of Ca 2+ stores by CPA we monitored intracellular Ca 2+ levels during drug treatment (0.05 Hz frame rate). Transient CPA-dependent Ca 2+ elevations lasted 10-40 min. After baseline Ca 2+ 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 mm slices were cut with a VT1000S vibratome (RRID:SCR_016495; Leica Biosystems, Nussloch, Germany). Acute slices were stored in a submerged, oxygenated storage container (S 2 ; 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 À1 collagenase A and 6 mg ml À1 DNase (10 min; 34˚C; shaking water bath (60 cycles min À1 )). 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 À1 trypsin and 20 mg ml À1 DNase (20 min; 34˚C; shaking water bath (60 cycles min À1 )). Digestion was stopped by addition of 100 mg ml À1 soybean trypsin inhibitor (SBTI) and 20 mg ml À1 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; 400 g) and the supernatant discarded. The pellet was resuspended in DMEM containing FBS (10%) and penicillin G/streptomycin (1%), filtered (cell strainer (100 mm)), and cells were plated in 75 cm 2 cell culture flask (T75; Invitrogen) and placed in a humidified incubator (37˚C; 5% CO 2 ). Approximately 1 / 3 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; 800 g) and the supernatant discarded. The pellet was resuspended in DMEM at cell densities of~10 5 cells ml À1 and plated again either in culture flasks or on glass coverslips in 35 mm dishes for experimental use. Again, 1 / 3 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:

Electrophysiology
Whole-cell patch-clamp recordings were performed as described (Fleck et al., 2016;Veitinger et al., 2011). 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 S 1 (~3 ml min À1 ; gravity flow;~23˚C). Patch pipettes (~5 MW) 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 S 6 . An agar bridge (150 mM KCl) connected reference electrode and bath solution. An EPC-10 amplifier (RRID:SCR_018399) controlled by Patchmaster 2.9 software (RRID:SCR_000034; HEKA Elektronik) was used for data acquisition. We monitored and compensated pipette and membrane capacitance (C mem ) as well as series resistance (R series ). C mem values served as a proxy for the cell surface area and, thus, for normalization of current amplitudes (i.e. current density). Cells displaying unstable R series 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 1 / 3 -1 / 5 of the sampling rate (10 kHz)]. If not stated otherwise, holding potential (V hold ) was -60 mV.

Fluorescence Ca 2+ imaging
Cultured mouse TPCs were imaged as described (Veitinger et al., 2011). Briefly, cells were loaded with fura-2/AM in the dark (5 mM; 30 min; RT; S 1 ) and imaged with an upright microscope (Leica DMI6000FS, Leica Microsystems) equipped for ratiometric live-cell imaging with a 150 W 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 f 340 /f 380 intensity ratio.
For parallel recordings of intracellular Ca 2+ signals and tubular contractions, acute seminiferous tubule slices were bulk-loaded with fura-2/AM in the dark (30 mM; 30 min; RT). After washing (3x; S 1 ), 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 Ca 2+ and tubular movement.
To ensure effective store depletion by CPA treatment, we recorded intracellular Ca 2+ levels during CPA incubation at low frequency to monitor Ca 2+ release from the ER, but also prevent phototoxicity. Experiments were only conducted if (i) we detected a substantial gradual rise in intracellular Ca 2+ upon CPA treatment, and if (ii) functional Ca 2+ extrusion mechanisms ensured that Ca 2+dependent fluorescence signals returned to base level. The time-course of this Ca 2+ release -Ca 2+ extrusion process varied between samples and ranged between 5.3 and 44.0 min (18.8 ± 9.3 min; mean ± SD).

Whole-mount seminiferous tubule imaging
Isolated tubules (>1 cm length) were placed onto a membrane within a custom-built 3D printed twocompartment recording chamber that was constantly superfused with S 1 . 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 S 1 from the top compartment. After visual determination of tubular stages (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 mM; 10 s) and corresponding negative controls were compared to determine ATP-dependent Ca 2+ signals (offspring from crossing SMMHC-CreER T2 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 À1 body weight) to double-positive adult male offspring (SMMHC-CreER T2 x Ai95D) via daily intraperitoneal injections for five 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, 10, 0.05-0.1 mg kg À1 , 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 (S 1 ; 35˚C), mounted on a 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. Individual recording duration varied between 13 and 30 min (mean = 25 min). For in vivo stimulation experiments, we used a Picospritzer III (Parker Hannifin, Pine Brook, NJ) to puff nanoliter volumes of control saline (S 1 ; containing Alexa Fluor 555 (4 mM)) or stimulus solution (S 1 ; containing Alexa Fluor 555 (4 mM) 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 (RRID:SCR_000325; WaveMetrics), Excel 2016 (Microsoft, Seattle, WA), and Leica LAS X (RRID:SCR_013673; Leica Microsystems) software. Dose-response curves were fitted by the Hill-equation. Time-lapse live-cell imaging data displaying both Ca 2+ signals and tubular contractions were analyzed using custom-written code in MATLAB (RRID:SCR_001622; 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 t 0 , 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 (f 340 + f 380 ) and then applied the displacement vector field, computed by the registration algorithm, to both images (f 340 and f 380 ) separately. ROIs were defined manually at t 0 and superimposed onto all subsequent images of the stabilized recording. At each time point t i , the fluorescence signal F was computed as the mean f 340 /f 380 ratio of all pixels within a given ROI. When measuring Ca 2+ -dependent changes in GCaMP6f intensity, the fluorescence signal F was normalized with respect to a baseline before stimulation, computing the intensity change for the i th time point as Fi À Fbaseline Fi . 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 t i , the registration algorithm computed a flow or displacement vector field V i ¼ X kv p;q k is a measure for the effort that is necessary to register the image at t 0 to the image at t i . 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 s i within an ROI as the average norm jV i | computed only for the v p;q corresponding to pixels within the ROI defined at t 0 . For whole-mount macroscopic imaging of luminal content movement in intact tubule segments ( Figure 5), we quantified the flow change c i ¼ s i À s iÀ1 as the change of flow strength between two consecutive time points / frames. Here, s i values were preprocessed by smoothing with a moving average filter. Results are reported as the AUC, that is, the area under the c i curve. For analysis of in vivo data, we employed a custom set of ImageJ macros utilizing build-in functions of Fiji-ImageJ (RRID:SCR_002285) (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 (five 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 mm 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 (five 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). Time-lapse epifluorescence in vivo recordings were processed to isolate transient fluorescence signals from static background noise using custom ImageJ code with Fiji's build-in functions (see Data and materials availability).

Ethics
Animal experimentation: 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).