We previously identified seven protein components of the CatSper channel complex (CatSper1-4, β, γ, and δ) from mouse testis using tandem affinity purification (Chung et al., 2011). As the most biochemically complex ion channel known to date, it has not been possible to express functional CatSper channels in heterologous systems. This includes many attempts in many cell types, including simultaneous injection of all 7 CatSper mRNAs into Xenopus oocytes (data not shown). Therefore, we continued to seek potential additional components to more thoroughly understand CatSper channel assembly and trafficking. We identified a mouse homolog of human C1orf101 (C1orf101-like, currently Gm7068) (Figure 1A) based on its sequence homology to the C-terminal extracellular domain of CatSperδ (Figure 1—figure supplement 1A). This testis-specific gene (Figure 1—figure supplement 2A) is predicted to encode a single transmembrane (TM) protein (Figure 1C and Figure 1—figure supplement 1B,C). In addition, a small soluble protein encoded by another testis-specific gene, Tex40 (Figure 1—figure supplement 2A), was found to be associated with the CatSper channel complex (Figure 1B and C, and Figure 1—figure supplement 1D). In this study, we refer to the C1orf101-like and Tex40 genes as CatSpere and CatSperz, respectively (see Molecular Cloning, Materials and methods). Like the other CatSper accessory subunits (Chung et al., 2011), both CatSpere and CatSperz mRNAs express specifically in germ cells and are detected before CatSper1 expression during postnatal development (Figure 1—figure supplement 2B,C). Moreover, mouse CatSper ε and ζ proteins partition into the testis microsome fraction (P) (Figure 1—figure supplement 2D), complex with CatSper1, and exhibit interdependence with the expression of the other CatSper subunits (Figure 1D–1F). In both human and mouse sperm cells, CatSper ε and ζ proteins are localized to the principal piece of the tails (Figure 1G and H and Figure 1—figure supplement 2E–G).

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Mouse CatSper proteins form a unique pattern of four linear (‘racing stripes’) Ca²⁺ signaling domains running down the four quadrants of the principal piece of the flagellum (Chung et al., 2014). We examined whether ε and ζ share this distinctive compartmentalization. The antibodies, anti-hε31, recognizing the N-terminal extracellular region of human CatSperε, and anti-mζ174, against the very C-terminus of mouse CatSperζ, were suitable for 3D stochastic optical reconstruction microscopy (STORM) (Figure 1F–1H and Figure 1—figure supplement 2E). CatSperζ and CatSperε show the apparent four-fold arrangement of CatSper1, β and δ subunits in mouse (Figure 1I) and human (Figure 1J) spermatozoa.

The lack of functional expression of CatSper channels in heterologous systems requires that genetic manipulation be used to study the function of each component. CatSpere has the same ancient origin at the root of early eukaryotic evolution as those of CatSpers1-4, b, and g and the same pattern of extensive lineage-specific gene loss as CatSperb and g through metazoan evolution (Figure 2—figure supplement 1A) (Cai et al., 2014). While CatSper δ and ε share high C-terminal sequence homology (Figure 1—figure supplement 1A), CatSperδ appears later in evolution (Figure 2—figure supplement 1A). In contrast, CatSperζ has no conserved domains and, like hyperactivated motility, is only present in mammals (Figure 2—figure supplement 1A), leading us to speculate that CatSperζ is a required evolutionary adaptation to mammalian fertilization. Based on sequence homology and conservation, we anticipated that deletion of CatSperε would likely be the same as the existing knockout of other CatSper subunits, but deletion of CatSperζ might provide new insights into spermatozoan adaptations to changes concomitant with the evolution of mammalian fertilization. To test this idea, we began by generating a CatSperz-null mouse line from Tex40 gene targeted ES cell clones. Tex40 is a small gene composed of 5 exons that spans only ~3 kb on chromosome 11 (Figure 2—figure supplement 1B). Deletion of exons 2–4 was confirmed in the homozygous null mouse (Figure 2—figure supplement 1C). No CatSperζ protein was detected in Tex40-null spermatozoa by immunoblotting and immunocytochemistry (Figure 2—figure supplement 1D,E).

CatSperz-null mutant mice are indistinguishable from their wt or heterozygous (het) littermates in appearance, gross behavior, or survival. In addition, no morphological differences were observed by histological examination of testis and epididymis (data not shown). Sperm morphology and epididymal sperm number from CatSperz-null mice were not significantly different from those of 2–3 month old paired heterozygous littermates (Figure 2—figure supplement 1E and Figure 2—figure supplement 2A). CatSperz-null female mice exhibited normal mating behavior and gave birth to litters comparable to those of het females when mated with wt or het males (Figure 2—figure supplement 2B). However, when CatSperz-null male mice were mated with wt or het females, they were severely subfertile: 20% (5/25) CatSperz-null males were completely infertile over six months (Figure 2A), and progeny of the fertile paternal CatSperz-null mice were significantly fewer in number (Figure 2B and Figure 2—figure supplement 2B). The latency from pair formation to the birth of these offspring from CatSperz-null males was ≥10 days compared to those from wt or het males (data not shown).

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Impaired male fertility in CatSperz-/- mice: pregnancy rate, litter size, sperm number per egg, and in vivo fertilization rate.

https://doi.org/10.7554/eLife.23082.008

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Inward CatSper current at −100 mV.

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We examined the number of sperm within cumulus oocyte complexes (COCs) after copulation and checked in vivo fertilization rates by isolating the COCs and/or embryos from the female ampullae. At 8 hr after coitus, no sperm was found in the COCs when mated with CatSperz-null male mice (Figure 2C). In contrast, the majority of the COCs from CatSperz-het mated females had one or more sperm cells within the complex. When mated with CatSperz-het males, more 2 cell eggs were observed over time after coitus, while the fertilization rate by CatSperz-null males did not change significantly (Figure 2D and Figure 2—figure supplement 2C). These data suggest that CatSperz-null sperm migration is delayed in the female reproductive tract.

To understand why CatSperz-null spermatozoa did not efficiently migrate in the female reproductive tract, we first investigated sperm motility using computer assisted sperm analysis (CASA) (Figure 2—figure supplement 2D). The percentage of motile spermatozoa was not significantly different and most motility parameters of z-null spermatozoa were comparable to those of z-het sperm cells. However, the characteristic increase of lateral head displacement upon capacitation was not observed in z-null spermatozoa (Figure 2—figure supplement 2D), suggesting that hyperactivated motility was reduced. Ninety minutes after capacitation, there was a less marked difference in swimming trajectories of z-null spermatozoa compared to z-het spermatozoa, supporting this notion (Figure 2E and Video 1). Further analysis of flagellar amplitude and waveforms of tethered spermatozoa revealed a striking rigidity of z-null spermatozoa from their midpiece to midway down the principal piece (Figure 2F and Video 2). This phenotype was also observed from hyperactivation-deficient CatSper2-null patients (Smith et al., 2013). After incubation under capacitating conditions for 90 min, we observed that z-null spermatozoa beat only at the very distal end of a flagellum (Video 3). Moreover, CatSperz-null spermatozoa remain bent in the anti-hook direction (Ishijima et al., 2002) (Figure 2F, Videos 2 and 3). The anti-hook bend predominates as the pro-hook bend (initiated by the CatSper-mediated Ca²⁺ signaling pathway (Chang and Suarez, 2011) is dysregulated in CatSperz-null spermatozoa.

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To examine how Ca²⁺ signaling in CatSperz-null spermatozoa is impaired, we first examined I_CatSper, the sperm-specific Ca²⁺-selective ion current. Since Ca²⁺ has high affinity to calcium-selective pores (Almers et al., 1984), CatSper permeation of monovalents increases when external calcium is removed (Kirichok et al., 2006; Navarro et al., 2007). In divalent-free (DVF) solutions, wt spermatozoan I_CatSper conducts a large Na⁺ current, which is completely absent in mice lacking CatSpers1, 2, 3, 4, or d (Chung et al., 2011; Kirichok et al., 2006; Qi et al., 2007). However, in CatSperz-null spermatozoa, monovalent CatSper current is present but reduced to ~60% of normal (−426 ± 50 pA at −100 mV; Figure 3A) compared to control CatSperz-het spermatozoa (−683 ± 77 pA at −100 mV; Figure 3B). Thus, in the absence of CatSperζ, the CatSper channel complex is still targeted to the flagellar membrane and forms functional channels. We hypothesize that the reduction in CatSper current reflected decreased protein expression levels.

P2X receptors are nonselective ion channels gated by purines such as ATP. The ATP-activated cation-nonselective current in the midpiece of murine sperm is mediated by the P₂X₂ purinergic receptor (Navarro et al., 2011). In CatSperz-null spermatozoa, I_ATP did not differ substantially from heterozygous spermatozoa (Figure 3A and B), supporting the assumption that there is selective down regulation of CatSper channels. Smaller I_CatSper explains, in part, the attenuated hyperactivated motility, delayed sperm migration, and male subfertility (Figure 2A–2E). Protein tyrosine phosphorylation (P-Tyr), a hallmark of sperm capacitation, is potentiated and delocalized in CatSper knockout mice (Chung et al., 2014) or when Ca²⁺ influx is pharmacologically blocked (Navarrete et al., 2015). Upon capacitation, P-Tyr was more prominent in CatSperz-null spermatozoa than wt, but to a lesser extent than CatSper1-null spermatozoa (Figure 2—figure supplement 2E), consistent with the reduced calcium current. It is, however, also possible that an altered arrangement of the CatSper complex and/or its interaction with target proteins in the linear domains could have contributed to these functional deficits.

To better understand why I_CatSper is reduced in CatSperz-null spermatozoa, we examined levels of protein expression in CatSperz-null spermatozoa (Figure 4A). Expression of other CatSper subunits was detected in CatSperz -null spermatozoa, albeit at 30–60% lower levels than that of wt (Figure 4B), consistent with reduced I_CatSper (Figure 3). This contrasts with the complete absence of other CatSper subunits in CatSper1- and CatSperd-null spermatozoa. mRNA and protein levels of other CatSper subunits were not reduced in the testis of CatSperz-null mice (Figure 4C and D), suggesting that the defect occurs during or after assembly of the protein complex.

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Protein and mRNA expression of CatSper subunits.

https://doi.org/10.7554/eLife.23082.018

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Loss of CatSperz resulted in fragmentation of CatSper1 staining on the flagellar membrane and these defects are large enough to be resolved by confocal imaging (Figure 5A). These gaps were not observed in wt/het (Figure 1H and Figure 2—figure supplement 1E) or previous wt and CatSper knockout studies (Chung et al., 2011; 2014; Liu et al., 2007; Ren et al., 2001). 3D STED and 3D STORM super-resolution microscopies clearly demonstrate that structural continuity is interrupted in CatSperz-null spermatozoa - each ‘stripe’ of the CatSper domains is fragmented, while the overall quadrilateral structure is maintained (Figure 5B and C). Cross-sections of the 3D STORM image of wt flagellum show the normal four tight clusters (Figure 5C, lower), represented as four lines in the 2D angular profiles of surface localizations (Figure 5—figure supplement 1A,E; inset) as previously observed (Chung et al., 2014). In CatSperz-null spermatozoa, however, the four lines in the 2D angular profiles were interrupted (Figure 5—figure supplement 1B). To examine whether the interruptions were periodic, we performed autocorrelation analysis and Fourier transform of STORM images of CatSperz-null sperm flagella (Figure 5—figure supplement 1C–F). Autocorrelation analysis of the CatSperz-null sperm flagella exhibited enhanced periodicity compared to the wt flagellum, with the first peak at ∼850 nm (Figure 5—figure supplement 1C,D). The Fourier transform shows a fundamental frequency of (800 nm)⁻¹ (Figure 5—figure supplement 1F). We assume this thinning of one or more linear domains reflects an underlying structural periodicity that regulates CatSper complex trafficking or membrane insertion.

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Thus far, our results show that CatSperz-null sperm have reduced I_CatSper, dysregulated structural continuity of the CatSper Ca²⁺ signaling domains, beat in an atypical pattern, and are delayed in migrating in the female reproductive tract, resulting in reduced male fertility. Rheotactic guidance for sperm over long distances requires rotational motion during CatSper-mediated hyperactivated motility (Miki and Clapham, 2013). We thus measured rheotactic parameters and the rotation rate of CatSperz-null spermatozoa with a particular focus on whether their proximal tail rigidity and subsequent low amplitude lateral movement (Figure 2) affects sperm movement. In flow-directed capillary tubes (Miki and Clapham, 2013), we observed that the rheotactic ability of CatSperz-null spermatozoa was significantly reduced (Figure 6A and B and Figure 6—figure supplement 1A). At all flow rates tested, most motile CatSperz-null spermatozoa were unable reorient to swim against the flow and were swept out of the tube (Video 4). In contrast, 85% of motile heterozygous spermatozoa displayed rheotactic behaviors by maintaining their position or swimming upstream for more than 2 s of the 9 s period of recording (Figure 6B and Video 4).

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In-capillary sperm rheotaxis and in vitro fertilization.

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We next examined the rotational motion of CatSperz-null spermatozoa. At high viscosities (0.3% methyl cellulose (MC), cP = 6.7), uncapacitated z-het spermatozoa swim in circles (Figure 6C, left and Video 5), while capacitated z-het spermatozoa swim in a more linear path as they rotate around a longitudinal axis, like wt spermatozoa (Video 6) (Miki and Clapham, 2013). Interestingly, z-null spermatozoa exhibit linear migration as they can rotate along the tail axis regardless of capacitating conditions, even at higher viscosities (Figure 6C, right and Videos 5 and 6). Indeed, uncapacitated CatSperz-null spermatozoa rotate ~50% faster than z-het spermatozoa (Figure 6D). This indicates that CatSperz-null spermatozoa have less lateral motion and are subject to less torque by the moving stream. Spatially, the spermatozoa trace out a less conical 3D envelope (Figure 2F and Figure 6—figure supplement 1B). In short, the rigidity of the CatSperz-null sperm proximal tail constrains its motion to that of a propeller-driven rod.

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We next examined the relation between external calcium entry and sperm function. First, we tested whether increasing extracellular [Ca²⁺] could rescue z-null sperm motility. After incubation for 90 min with a two-fold greater [Ca²⁺], most CatSperz-null sperm remain bent in the anti-hook orientation with a rigid proximal tail (Figure 6—figure supplement 1C, z(-/-) middle, and Video 7). A few z-null spermatozoa partially recover, bending occasionally in the pro-hook direction with hyperactivated motility (Figure 6—figure supplement 1C, z(-/-) right, and Video 7). Conversely, a 20-fold reduction of extracellular [Ca²⁺] alone did not significantly alter the flagellar waveforms of z-het spermatozoa within 90 min (Figure 6—figure supplement 1C and Video 8).

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A transient Ca²⁺ pulse induced by Ca²⁺ ionophore, A23187, significantly reduces the time required for wt sperm to develop hyperactivated motility (Tateno et al., 2013). Moreover, a short (10 min) exposure to A23187 can rescue defects in hyperactivated motility and the fertilizing capability of CatSper1-null sperm in vitro (Navarrete et al., 2016). We tested the relation of calcium transients to hyperactivated motility in CatSperz-het and null sperm. In z-het spermatozoa, an A23187-induced Ca²⁺ pulse followed by washout, enables full hyperactivation, characterized by wide lateral displacement with large midpiece α angle within 30 min (Figure 6—figure supplement 1D and Videos 9 and 10). However, in z-null sperm, the same treatment improved the flexibility of the proximal flagella, particularly in the principal piece, but the midpiece remained largely inflexible (Figure 6—figure supplement 1D and Videos 9 and 10). Building on our previous work, the present study suggests that calcium entry through CatSper channels has time-dependent, complex effects on the coordination of motility and that loss of CatSperz results in reduced I_CatSper, changes in calcium signaling, and structural alterations of the flagellum.

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We performed in vitro fertilization (IVF) to determine how the low rotational torque generated by CatSperz-null spermatozoa affects sperm-egg interactions. We found that these spermatozoa cannot fertilize cumulus-intact oocytes (Figure 6E), but could dissociate the cumulus cell layers and bind to the ZP (data not shown). Cumulus removal did not change the fertilization rate of ZP intact oocytes by CatSperz-null spermatozoa. Furthermore, this rate was only marginally enhanced by destabilization of the ZP by 2 mM glutathione (Figure 6F) (Miyata et al., 2015). This indicates that reduced hyperactivated motility alone does not explain the failure of CatSperz-null spermatozoa in IVF. One possibility is that the kinetics of capacitation in vitro is different from that in vivo, resulting from fluctuations in timing or amplitude of known factors (e.g., HCO₃, pH) or from unknown factors present in seminal and/or female fluids. We then compared IVF rates with ejaculated and epididymal spermatozoa of CatSperz-null mice. When ejaculated spermatozoa flushed from the uterus of the mated females were used in IVF trials, 20% of oocytes incubated with z-null spermatozoa developed into two-cell embryos (Figure 6E). This compares to 50% of oocytes incubated with z-het ejaculated (data not shown), or epididymal sperm. Thus, additional factors may be functionally relevant in in vivo fertilization.

Sperm hyperactivation and normal fertility in mammals requires the unique CatSper channel complex. With four distinct pore-forming gene products (CatSper 1–4) and, now, five accessory subunits (β, γ, δ, ε, and ζ), the CatSper channel is the most complex of known ion channels. This may reflect the relatively high evolutionary pressure on spermatozoan evolution (Swanson and Vacquier, 2002; Torgerson et al., 2002), and various adaptations to different modes of fertilization. Like many gamete-specific proteins and the other CatSper proteins reported so far (Cai and Clapham, 2008; Chung et al., 2011), mouse and human CatSper ε and ζ show signs of rapid evolutionary change with only 50% and 45% amino acid sequence identity, respectively. In particular, the sequence regions outside TM segments and the pore loop of CatSper proteins are poorly conserved across species, indicating these regions possibly convey species-specific modulation of flagellar motility (Miller et al., 2015). This is illustrated by striking differences in progesterone-elicited I_CatSper responses in mouse and human (Lishko et al., 2011). Here we have shown that CatSper ε and ζ are components of the highly organized CatSper complex, that CatSperζ is required for proper continuity of this complex along the flagellum, and that loss of ζ alters hyperactivation waveforms and reduces fertilizing capacity.

The conservation pattern of the lineage-specific gain and loss of the CatSpere gene is identical to those of b and g, suggesting that they likely belonged to an ancient CatSper channel Ca²⁺ signaling network before the divergence of unikonts and bikonts. Since their protein expression is strictly interdependent, we speculate that CatSpere-null mice will have the phenotype of CatSper1-4, or d-null mice. In contrast, CatSperz is conserved only in mammals, suggesting that this protein imparts some adaptation, perhaps as a method enabling rheotaxis in the mammalian female reproductive tract.

Interestingly, although CatSperζ has no putative transmembrane domains, it is localized in the same quadrilateral pattern as other CatSpers, but is not present elsewhere in sperm. An intriguing aspect of our observations is that, unlike CatSper1-4 and d-null mice, which produce complete infertility, CatSperz-null males exhibit an incomplete loss of fertility. The CatSper current is reduced in CatSperz-null spermatozoa, and may have similar permeation properties (likely dominated by the CatSper1-4 pore subunits), but the effects of CatSperz on channel gating remain to be determined in future studies. This is reminiscent of the non-spermatozoan, voltage-gated Ca_v channel auxiliary subunits, which are not required for expression but modulate expression levels and gating (Catterall et al., 2005). Most tantalizing is the thinning and disruption of the linear CatSper signaling domains at repeat intervals in the absence of ζ. Further detailed examination via mutagenesis experiments has been stymied by our inability to heterologously express functional CatSper channels. New rapid genome editing techniques should enable more mice to be generated that will further the study of CatSper trafficking, subunit interactions, and localized signaling pathways.

Functional Ca²⁺ signaling domains are common adaptations in many biological systems, such as synapses and muscle. They enable specific and fast triggering of downstream events (Clapham, 2007). CatSper channels are compartmentalized into a unique multilinear arrangement and form Ca²⁺ signaling nanodomains with other Ca²⁺ signaling molecules along the sperm flagellum (Chung et al., 2014). The mechanisms involved in the delivery of the CatSper channels to these specific domains are currently unknown, and we suspect will be as interesting and complex as those in primary and motile cilia (Sung and Leroux, 2013). We found that abrogation of CatSperz not only retards targeting of the CatSper complex to flagella, but also disrupts continuity of the linear domains, resulting in repeated fragmented domains with ~800 nm periodicity. In order for CatSper domains to form and function properly, interactions are needed between the CatSper channel complex in the flagellar membrane and the underlying cytoskeletal proteins. One speculation is that CatSperζ might adapt to cytoskeletal structures that traffic, distribute, and enable membrane insertion of CatSper.

The fibrous sheath (FS), a cytoskeletal structure unique to the mammalian sperm flagellum, defines the extent of the tail’s principal piece, in which all the CatSper proteins are specifically localized. The FS closely lies under the plasma membrane and its two longitudinal columns are connected by circumferential ribs. Immunogold electron microscopy demonstrated that the CatSper channels are distributed on the end of ribs, where they merge with the column (Chung et al., 2014). It seems likely that the timing of occurrence and localization of CatSper Ca²⁺ signaling domains is coordinated with the assembly of FS proteins along the axoneme. The column appears early in spermiogenesis, forming from the distal tip of the tail along the axoneme, followed by subsequent rib formation in the opposite direction (Oko, 1998; Oko and Clermont, 1989). Based on scanning electron micrographs (Danshina et al., 2010; Miki et al., 2004), we find that the distance between ribs is about 800 nm in mouse spermatozoa. Thus, it seems likely that the repeated disruption in the absence of CatSperz is related to rib spacing of the FS.

Genetic abrogation of CatSper disrupts hyperactivated motility as manifested by changes in movement symmetry, amplitude, and rolling (Carlson et al., 2003; Chung et al., 2011; Miki and Clapham, 2013; Qi et al., 2007). Here we report that the flagellar envelope is significantly altered in the absence of CatSperz, in part due to the inflexibility of the proximal tail. We previously reported that the catalytic subunit of calcineurin, PP2B-Aγ, expresses throughout the tail but localized to the CatSper quadrilateral structures and axoneme (Chung et al., 2014). In CatSper1-null spermatozoa, PP2B-Aγ remains localized primarily to the axoneme but disappears from the quadrilateral structures. Recently, a similar but not identical phenotype (inflexible midpiece, reduced hyperactivated motility, and impaired ZP penetration) was reported in testis-specific calcineurin Ppp3cc-null and Ppp3r2-null spermatozoa (Miyata et al., 2015). Note that the principal piece of both CatSper1-null and Ppp3cc-null spermatozoa are not rigid. The integrity and distribution of CatSper channels in Ppp3cc-null spermatozoa remain to be examined and may clarify midpiece/principal piece disparities. In any case, inflexibility in the proximal regions of flagellum results in a flagellar envelope approximated as a rod with a distal propeller. The sperm can rotate faster but the smaller lateral deviation reduces torque. This limits the sperm’s ability to orient into the flow, as well as penetrate the cumulus and ZP.

Gene-manipulated mice highlight the importance of in vivo observations and have reshaped the landscape of fertilization science (Okabe, 2015). In vitro capacitation and fertilization systems underpin much of the study of sperm motility and fertilization potential. While ejaculated sperm are preferred for fertilization studies in larger animals and humans, epididymal sperm are commonly used in genetically tractable mouse studies. Notably, these sperm are not exposed to accessory sex gland secretions and female fluids. This may explain why CatSperz-null spermatozoa are completely infertile in an IVF setting (COCs), but in vivo are merely subfertile. Perhaps natural modulators, absent in epididymal sperm IVF studies, partially rescue the fertilizing potential of CatSperz-null spermatozoa by activating Ca²⁺ signaling activity.

Interestingly, a transient pulse of Ca²⁺ can greatly reduce the capacitation time required for wt sperm to develop hyperactivated motility (Tateno et al., 2013). Moreover, Navarrete et al recently demonstrated that a short exposure to A23187 rescued the defects in motility and fertilizing capability of CatSper1-null sperm in vitro (Navarrete et al., 2016). These independent studies were interpreted to mean that the initial priming by Ca²⁺ influx, perhaps above a certain threshold, is essential for sperm function. However, the linear quadrilateral CatSper complexes are not present in CatSper1-null spermatozoa and in CatSperz-null spermatozoa are disrupted by gaps. We hypothesize that the linear quadrilateral structure in vivo likely maintains, regulates, and distributes CatSper Ca²⁺ signaling during hyperactivated motility. But it is important to point out that alterations in the structure should also result in changes in mechanical properties, movement of the flagellum, distribution of entering calcium, and downstream kinase activity and the motor elements they regulate. This complexity is illustrated in vivo sperm swimming trajectories, which are modulated by switching between pro- and anti-hook beating patterns. In the absence of CatSperζ, anti-hook beating predominates. Pro-hook motions are associated with intact CatSper-mediated Ca²⁺ signaling pathways (Chang and Suarez, 2011). Finally, ejaculated sperm display more pro-hook hyperactivation than epididymal sperm (Li et al., 2015).

Future areas for investigation are the functional positioning of the remaining accessory subunits of the CatSper channel in assembly and domain organization, the testing of potential modifiers present in accessory sex gland secretions that may activate CatSper channels, and the determination of Ca²⁺ dependent molecules in the axoneme which eventually determine flagellar bending and its envelope. CatSperz-null mice, which are hypomorphic to the null-mutation of other CatSper genes with abrogated hyperactivation, and newly expanding animal models from recent advances in genome editing will serve as a foundation to this end. Advanced imaging techniques with higher time and spatial resolution will be necessary to carry this out. The present results also suggest that alterations of Ca²⁺ current and/or dysregulated downstream Ca²⁺ signaling affecting dynamic structures may be sufficient to compromise sperm function. CatSper’s unique composition and central role in hyperactivated motility make it an ideal target for contraception.

CatSper1 and d-null mice were previously described (Chung et al., 2011; Ren et al., 2001). Lines were backcrossed and maintained on a C57BL/6 background. WT C57BL/6 male, B6D2F1 female (Jackson laboratory, Bar Harbor, ME), and CD1 (Charles River Laboratories, Wilmington, MA) female mice were purchased.

Mouse caudal epididymal sperm were collected by swim-out in HEPES buffered saline (HS) containing (in mM): 135 NaCl, 5 KCl, 2 CaCl₂, 1 MgSO₄, 20 HEPES, 5 glucose, 10 lactic acid, 1 Na pyruvate, pH 7.4 (with NaOH) (Chung et al., 2011). To induce capacitation in vitro, sperm cells were incubated (2 × 10⁶ cells ml⁻¹) in human tubular fluid (HTF) media (in mM): 102 NaCl, 4.7 KCl, 2 CaCl₂, 0.2 MgCl₂, 0.37 KH₂PO₄, 2.78 glucose, 18.3 lactic acid, 0.33 Na pyruvate, 25 HCO₃^- and 4 mg ml⁻¹ BSA) (Millipore) for 90 min at 37°C (5% CO₂).

All experiments using human samples were approved by the Committee of Clinical Investigation, Boston Children’s Hospital CCI/IRB (IRB-P00000538). Human semen samples were obtained from fertile donors. Human spermatozoa were collected by the swim-up method with the use of modified human tubal fluid medium (HTF).

HEK293T cells were purchased from ATCC. In this study, they were used to overexpress recombinant human CatSperζ in order to test antibodies. The cells were tested negative for mycoplasma and validated as of human origin. The identity was authenticated by confirming their negative expression of testis-specific genes including CatSper. The cell line was cultured in DMEM/F12 containing 10% FBS.

Rabbit polyclonal CatSper1, CatSper4, β, and δ antibodies were previously described (Chung et al., 2011; Ren et al., 2001). To produce antibodies to new CatSper subunits, peptides were synthesized and conjugated to KLH carrier protein (Open Biosystems, Lafayette, CO) as follows: mouse CatSperε, 968–985 (αm-ε968: RQFIIEPLHKRPAKQKKN); mouse CatSperζ, 174–195 (αm-ε174: GYIEGIRKRRNKRLYFLDQ); human CatSperε, 31–50 (αh-ε31: RIFSTRSTIKLEYEGTLFTE); and human CatSperζ, 11–29 (αh-ζ11: KSSDRQGSDEESVHSDTRD). Antisera were affinity purified on the immobilized resin of the corresponding peptide (Amino Link Plus or Sulfo Link Plus) (Pierce, Waltham, MA). Anti-phosphotyrosine (clone 4G10), anti-Flag (clone M2), anti-calmodulin (05-173) and anti-acetylated tubulin (T7451) antibodies were from EMD Millipore (Germany). All chemical compounds were from Sigma-Aldrich (St. Louis, MO) unless indicated.

Annotated orthologs in the NCBI gene database (http://www.ncbi.nlm.nih.gov/gene/) and/or homologous amino acid sequences of reported protein databases were screened in 17 eukaryotes for the presence of genes for CatSper auxiliary subunits. Non-annotated orthologs in the NCBI gene database were identified by comparing sequences of the annotated orthologs to those in the protein database of species by Phmmer implemented on HMMER 3.1 (default option, http://hmmer.org/). The longest amino acid sequences among all the isoforms of the orthologs annotated in each species and protein sequence databases from 15 eukaryotes, except human and mouse, were downloaded from the NCBI Genome database (http://www.ncbi.nlm.nih.gov/genome; Tinamus guttatus, GCA000705375.2; Anolis carolinensis, GCA000090745.2; Salmo salar, GCA000233375.4; Callorhinchus milii, GCA000165045.2; Branchiostoma floridae, GCA000003815.1; Caenorhabditis elegans, GCA000002985.3; Crassostrea gigas, GCA000297895.1; Exaiptasia pallida, GCA001417965.1; Trichoplax adhaerens; GCA000150275.1, Salpingoeca rosetta, GCA_000188695.1), Ensembl genome browser (http://ensembl.org; Strongylocentrotus purpuratus, GCA000002235.2; Drosophila melanogaster, GCA000001215.4; Thecamonas trahens, GCA000142905.1), and JGI genome portal (http://genome.jgi.doe.gov; Allomyces macrogynus; Aurantiochytrium limacinum). Aligned phmmer hits of expected values <10⁻¹⁰ were considered as candidate orthologs of the corresponding CatSper subunits in each species.

PCR was performed according to standard protocols using a commercial multiple panel cDNA template (MTC), Clontech). PCR primers amplified Gm7068 (forward: 5′-CTATGGCTCAAGTGTAATGACC-3′, reverse: 5′-GCTCTTATTGAATCCTCGAACC-3′), Tex40 (forward: 5′-GAAACAGGATTCGCAAGTACAG-3′, reverse: 5′-TCGTGGACCTATATGTGATGAG-3′) using mouse GAPDH (forward: 5′-TGAAGGTCGGTGTGAACGGATTTGGC-3′, 5′-ATGTAGGCCATGAGGTCCACCAC-3′) as a control.

The initial mouse Tex40 cDNA sequence (NM_001039494) was identified from database searches using novel peptide sequences from MS. The full-length human Tex40 cDNAs was obtained by PCR with primers (forward: 5′-GGGCAGAACCATGGAGGAAA-3′, reverse: 5′-AGGACTCAAATTCCACTCGGATG-3′) using the human testis cDNA library (Clontech). Sequencing the TOPO-cloned PCR products into pCR4-TA (Invitrogen) confirmed the full-length human Tex40 ORF, which was subcloned into pCMV-Tag2A (Stratagene) to express recombinant N-terminal Flag-tagged human CatSperζ in mammalian cells. Mouse Gm7068 was identified by homologous amino acid sequence to C-terminal CatSperd (Tmem146). There are six transcript variants (Almers et al., 1984); XM_006497083, 2; XM_006497084, 3; XM_017314031, 5: XM_006497085, 6; XM_017314033, and 8; XM_006497087). Variants 1, 3, 5, 6, and 8 are predicted to encode polypeptides with the same C-terminal sequence that can be detected by anti-mε−968. Among them, the predicted polypeptides from longer splicing variant 1 (isoform X1; XP_006497147, 985 aa) and variant 3 (isoform X3; XP_017169520, 914 aa) are consistent with the apparent molecular weight of the band observed in testes microsomes (Figure 4C and Figure 1—figure supplement 2D). The predicted polypeptides from shorter variant 5 (isoform 4; XP_006497148, 805 aa) and variant 6 (isoform 5; XP_017169522, 770 aa) are consistent with that of the band detected in CatSper1-IP from testis and total sperm lysate (Figures 1D,E and and 4A). It is likely that mouse Gm7068 expresses at least four potential splice variants that can encode protein isoforms and/or undergo cleavage during spermatogenesis.

In situ hybridization experiments were carried out with an RNAscope (Advanced Cell Diagnostics, Newark, CA). Testes from three month old wild-type mice were fixed in 10% (vol/vol) neutral-buffered formalin at room temperature for 24 hr, dehydrated, and embedded in paraffin. Paraffin sections (10 μm thick) were processed according to the manufacturer’s instructions for in situ detection in the Rodent Histopathology Core Facility at Harvard Medical School. Sequences of the probes used in this study are: Gm7068 (XM_982472.3, 645–1072) and Tex40 (NM_001039494.2, 41–456). After the DAB (3,3, -diaminobenzidine) reaction, slides were counterstained using hematoxylin.

Real-time PCR was carried out with first strand cDNAs (iScript cDNA Synthesis) (Bio-Rad, Hercules, CA) synthesized from 2 µg total mouse testis RNA using the SYBR Green (iTaq Universal SYBR Green Supermix) (Bio-Rad; CFX96). Quantitative analysis by the ddCt method employed c-Jun as an amplification control. Three independent sets of experiments were performed to calculate fold changes (2^-ddCt) of CatSpers mRNA. The primers used for qRT-PCR were: CatSper1 (forward: 5′-CTGCCTCTTCCTCTTCTCTG-3′, reverse: 5′-TGTCTATGTAGATGAGGGACCA-3′), CatSperb (forward: 5′-CCTTA TTGACCAAGAAACAGAC-3′, reverse: 5′-TGAAACCCATATTTGACTGCC-3′), CatSperg (forward: 5′-TGAGCAATAGAGGTGTAGAC-3′, reverse: 5′-CAGGA TGTAGAAGACAACCAG-3′), CatSperd (forward: 5′-GCTGACATTTCTGTGTATCTAGG-3′, reverse: 5′-CTGATATACCTTCCAATTTACGCC-3′), CatSpere (forward: 5′-GTCTCATGCTTCTTCAGTTCC-3′, reverse: 5′- CAGAAGTTCCTTGTCCATCAC-3′), CatSperz (forward: 5′-GAGACCTCCTTAGCATCGTC-3′, reverse: 5′-TCGTGGACCTATATGTGATGAG-3′ and c-Jun (forward: 5′-CTCCAGACGGCAGTGCTT-3′, reverse: 5′-GAGTGCTAGCGGAGTCTTAACC-3′).

Testes (200 mg, normally two testicles) from 8- to 12-wk-old male mice were homogenized on ice using a Dounce homogenizer in 2 mL 0.32 M sucrose solution with protease inhibitor cocktails (Roche). The tissue suspension was centrifuged at 300 g for 10 min at 4°C and the supernatant was then transferred to an ultra-speed centrifuge tube. The microsome faction was isolated by centrifuging the tube at 105,000 g for 60 min.

Mouse sperm total protein was prepared as described before (Chung et al., 2011; 2014). For total protein from human spermatozoa, purified swim-up sperm were then lysed (0.1% SDS, 0.5% sodium deoxycholate, 1 mM DTT, 1 mM EDTA in PBS with protease inhibitors) followed by sonication for 5 min and centrifuged at 15,000 g for 10 min. The supernatants were further denatured by adding DTT to 10 mM and heated at 75°C for 10 min before SDS-PAGE. For immunoprecipitation, the testis microsome pellet was resuspended in 10 mL 1% Triton X-100 in PBS with protease inhibitors (Roche). The suspension was rocked at 4°C for 1 hr and then centrifuged at 15,000 g for 30 min. 1.5 mL of the solubilized testis microsome were mixed with 1–2 μg antibody and 25 μL Protein A/G-bead slurry (Santa Cruz Biotechnology) at 4°C overnight. The IP products were finally eluted in 50 μL LDS loading buffer containing 50 mM DTT. Antibodies used for Western blotting were rabbit anti-mouse CatSperε (αm-ε968; 1.6 μg/mL), mouse CatSperζ (αm-ε174; 2.7 μg/mL), human CatSperε (αh-ε31; 2.7 μg/mL), human CatSperζ (αh-ζ11: 1 μg/mL). Monoclonal anti-phosphotyrosine (clone 4G10; 1 μg/mL), anti-Flag (clone M2; 1 μg/mL), anti-calmodulin (05–173, 1 μg/mL), and anti-acetylated tubulin (T7451,1: 20,000). Secondary antibodies were anti-rabbit IgG-HRP (1:10,000) and anti-mouse IgG-HRP (1: 10,000) from Jackson ImmunoResearch (West Grove, PA).

Caudal epididymal mouse sperm cells attached to glass coverslips were fixed in 4% paraformaldehyde (PFA) in PBS, permeabilized with 0.1% TrixonX-100 for 10 min. Human sperm cells from swim-up purification were fixed 4% PFA in PBS for 10 min followed by 100% MeOH. Fixed human sperm cells were permeabilized in 0.1% saponin for 10 min. Permeabilized sperm cells were washed in PBS and blocked with 10% goat serum for 1 hr. Mouse samples were stained overnight with primary antibody against CatSper1 (10 µg ml⁻¹) and CatSperζ (mζ174, 20 µg ml⁻¹) as were human samples with primary antibodies against CatSperε (hε31, 20 µg ml⁻¹) and CatSperζ (hζ11, 10 µg ml⁻¹), in 10% goat serum in PBS, 4°C. After PBS wash, goat-anti-rabbit Alexa488 conjugate (Invitrogen) served as the secondary antibody. Images were acquired on laser scanning confocal microscopes (Olympus Fluoview 1000; Figure 1G and H, Figure 1—figure supplement 2G, and Figure 2—figure supplement 1E and Leica TCS SP8; deconvolved image in Figure 5A).

For timed coitus, females were introduced to single-caged CatSperz-het or -null males for 1 hr and checked for the presence of a vaginal plug. To examine sperm migration to the fertilization site in vivo, ampullae were removed from the mated females at 8 hr after coitus and COCs were released. A series of z-stacked images (2 µm step size) of the COCs was taken and number of sperm within each COC was recorded according to the presence of a sperm head. To calculate in vivo fertilization rate, eggs were gently flushed from oviducts and ampullae from the mated females at 20 hr and 27–30 hr after coitus. The total number of eggs and the number of 2 cell eggs were counted.

Two females were caged with each male for three months to track pregnancy and litter production. For IVF assays, oocytes were recovered from superovulated 5–6-week-old B6D2F1 female mice 13 hr after injection of 5 U human chorionic gonadotropin. For standard IVF, sperm were collected from the cauda epididymis. For ejaculate IVF, sperm were retrieved from the uterus of a 1 hr window-timed coitus. Both epididymal and ejaculated sperm were capacitated in vitro at 37°C for 1 hr, and coincubated with eggs at ∼10⁵ sperm/mL. After about 4.5 hr, unbound sperm were washed away. After 24 hr incubation the embryos were observed under light microscopy (Olympus IX-70) to check for development of the two-cell stage.

Spermatozoa from the dissected cauda epididymis (swim up method) were collected in HEPES buffered saline (HS) media. Spermatozoa were plated on 35 mm fibronectin-coated coverslips for 15 min (22°C); unattached sperm were removed by the gentle pipette wash (time 0) and basal motility recorded. Activated motility was recorded within the first 10 min after adding pre-warmed human tubal fluid (HTF)-capacitating medium (Millipore). To induce hyperactivation, attached sperm cells were incubated in HTF media for 90 min at 37°C (5% CO₂). All subsequent images were recorded at 37°C. The flagellar waveform was analyzed by stop-motion digital imaging collected at 200 fps (HC Image software, Hamamatsu Photonics or Zen Blue, Zeiss; 2 s movies). Overlay of flagellar traces from two complete flagellar beats were generated by hyperstacking binary images using open-source FIJI software (Schindelin et al., 2012) and time coded in color.

Cauda epididymal spermatozoa were suspended and incubated in non-capacitating M2 medium (Specialty Media, Millipore) or in HTF medium for capacitation. Sperm motility was then measured using the IVOS sperm analysis system (Hamilton Thorne Biosciences, Beverly, MA) in an 80 µm (depth) chamber to obtain various parameters (Figure 2—figure supplement 2D). Sperm motility was also analyzed with an Olympus IX-70 microscope equipped with a high-speed sCMOS camera (Orca-Flash4.0) and a 10x objective. 1–2 × 10⁵ mouse sperm before and after capacitation were added to the 37°C chamber (Delta T culture dish controller; Bioptechs) containing 1 ml HEPES-HTF medium (H-HTF: 92 mM NaCl, 2 mM CaCl₂, 4.7 mM KCl, 0.2 mM MgCl₂, 0.37 mM KH₂PO₄, 25 mM NaHCO₃, 18.3 mM Na lactate, 2.78 mM glucose, 0.33 mM Na pyruvate, 0.4% [w/v] bovine serum albumin [BSA], and 10 mM HEPES [pH 7.4]). In some experiments, the medium was supplemented with methylcellulose (MC) (M0512, 4000 cP in 2% solution; Sigma) at 0.3%, 0.4%, or 0.5% (w/v). Sperm swimming 3–5 mm from the rim were recorded after a 10 min preincubation period that allowed spontaneous dissociation of sperm clumps. To inhibit convective flow, 1 ml of medium was overlaid by 1 ml of mineral oil and covered by a heated glass lid (Bioptechs). Sperm motility at 37°C was videotaped at 100 fps. Images (HC Image software, Hamamatsu Photonics) were analyzed for swimming trajectory from a 1 s playback movie at 1/5 speed, by head tracing via Computer Assisted Sperm Analysis (CASA; http://rsbweb.nih.gov/ij/plugins/casa.html). To track swimming trajectory in viscous medium, the sperm motility was videotaped at 50 fps. The images were analyzed using Fiji software (Schindelin et al., 2012) by assembling overlays of the flagellar traces generated by hyperstacking binary images of 20 frames of 2 s movies coded in a gray intensity scale.

Mouse sperm incubated in HTF medium for 90 min at 2 × 10⁶/ml yielded capacitated sperm. Capacitated sperm were transferred and concentrated for capillary loading by centrifugation at 900 g for 3 min. The loose sperm pellet at the bottom of the microcentrifuge tube was resuspended in HEPES-HTF at 4 × 10⁶/ml, and loaded into the capillary by suction via an air-pressure microinjector (IM-5B; Narishige; 22 C, ~200 µm/s). While applying gentle positive pressure, the sperm in the tip of the capillary were moved out of the sperm drop. The tip of the capillary is transferred to a 37°C chamber (Delta T culture dish controller; Bioptechs) and placed into a 50 µl drop of HEPES-HTF medium covered with mineral oil. Negative pressure was applied slowly and sperm cells swimming against the flow and down to the H-HTF drop was video-recorded at 33 fps.

Whole-cell recording of corpus epididymal spermatozoa from 3–5 month-old CatSperz+/- or CatSperz-/- mice was performed blind as to genotype (Kirichok et al., 2006; Navarro et al., 2011). HS was the bath medium. The standard pipette solution was (mM): 120 Cs- Methanesulfonate (Cs-MeSO₄), 5 CsCl, 5 Cs-BAPTA, 10 HEPES and 10 MES, pH 7.2 with H-MeSO₃. To record I_ATP, we used a low Cl^- bath solution (to reduce background Cl^- conductance) in the following (mM): 150 Na- methanesulfonate (Na-MeSO₃), 2 CaCl₂, 10 Na-HEPES, and 10 MES (pH 7.4 or 6.0). To measure I_CatSper, we used divalent-free (DVF) solution, in mM: 150 Na-MeSO₃, 2 Na₃HEDTA [(hydroxyethyl)ethylenediaminetriacetic acid], 2 EGTA, and 20 HEPES (pH 7.4) with NaOH. Solutions were applied to sperm cells (lifted from the coverslips) initially by bath perfusion. After break-in, the access resistance was 25–80 MΩ. All experiments were performed at 22–24°C. The whole-cell currents were recorded using an Axopatch 200B amplifier (Molecular Devices, Sunnyvale, CA), acquired with Clampex 9 (pClamp9 Software; Molecular Devices), and analyzed with Origin software (OriginLab). Signals were low-pass filtered at 2 kHz and sampled at 10 kHz. Data are given as mean ± SD.

All the experiments are repeated at least three times. Sample size and number of replicates are described in each figure and the figure legends. Statistical analyses were performed using Student’s t-test unless indicated; e.g. F-test in one-way ANOVA. Differences were considered significant at *p<0.05, **p<0.01, ***p<0.001, and ****p<0.0001. When ****p<0.0001, actual P value is not indicated.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank M. Sanguinetti for testing CatSper expression in Xenopus oocytes for I_CatSper detection, P. DeCaen for help with sperm electrophysiology, and L. Ded for calculating rib length from the literature and critical reading of the manuscript-in-progress. This work was supported by Goodman-Gilman Yale Scholar Award 2015–08 (J-JC), HHMI (XZ), and HHMI (DEC).

Animal experimentation: This study was performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All the mice were treated in accordance with guidelines approved by the Boston Children's Hospital (13-01-2341R) and Yale (2015-20079) Animal Care and Use Committees (IACUC).

1. Received: November 8, 2016
2. Accepted: February 20, 2017
3. Accepted Manuscript published: February 22, 2017 (version 1)
4. Accepted Manuscript updated: February 23, 2017 (version 2)
5. Version of Record published: March 22, 2017 (version 3)

© 2017, Chung et al.

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