Sperm chromatin condensation: epigenetic mechanisms to compact the genome and spatiotemporal regulation from inside and outside the nucleus

ABSTRACT

Sperm chromatin condensation is a critical step in mammalian spermatogenesis to protect the paternal DNA from external damaging factors and to acquire fertility. During chromatin condensation, various events proceed in a chronological order, independently or in sequence, interacting with each other both inside and outside the nucleus to support the dramatic chromatin changes. Among these events, histone–protamine replacement, which is concomitant with acrosome biogenesis and cytoskeletal alteration, is the most critical step associated with nuclear elongation. Failures of not only intranuclear events but also extra-nuclear events severely affect sperm shape and chromatin state and are subsequently linked to infertility. This review focuses on nuclear and non-nuclear factors that affect sperm chromatin condensation and its effects, and further discusses the possible utility of sperm chromatin for clinical applications.

INTRODUCTION

During the past decade, evidence has accumulated for transgenerational epigenetic inheritance, sometimes referred to as “Lamarck revisited”. Importantly, not only in non-mammalian organisms but also in the rodent model and human cohort studies, events that strongly support the inheritance of acquired traits have been reported consistently (Bošković and Rando, 2018; Liberman et al., 2019; Duempelmann et al., 2020; Le Blévec et al., 2020; Senaldi and Smith-Raska, 2020). In 2012, John Gurdon and Shinya Yamanaka won the Nobel Prize in Physiology or Medicine for their work on “the discovery that mature cells can be reprogrammed to become pluripotent” (Gurdon, 1962; Gurdon et al., 1975; Takahashi and Yamanaka, 2006; Takahashi et al., 2007). Their work has accelerated the study of mechanisms for epigenetic reprogramming, especially regarding how to erase and re-establish epigenetic information. The transgenerational effect of the epigenome is, on the other hand, the question of what remains of the gametes’ epigenomic information that is not erased (Meyer et al., 2017).

The details of gamete epigenetic information, however, remain largely unexplored compared to those of somatic cells. One reason for this is the difficulty of in vitro culture and differentiation. In the last decade, this problem has been successfully solved in oocytes through intensive efforts utilizing iPS/ES cell technology (Hayashi et al., 2012; Hikabe et al., 2016; Hamazaki et al., 2021). For in vitro spermatogenesis, similar iPS/ES cell technology may achieve the generation of spermatogonial stem cells; in fact, an efficient ex vivo culture system of seminiferous tubules has been established (Hayashi et al., 2011; Sato et al., 2011). However, in vitro spermatogenesis is still under investigation, even though much effort has been expended over the past few decades (Makar and Sasaki, 2020; Pelzman et al., 2020). One possible reason for this is the inefficiency of meiotic progression in vitro owing to impaired chromosome synapsis and meiotic crossover (Lei et al., 2020). In the past five years, single-cell RNA-sequencing (scRNA-seq) analysis has been used to great advantage in spermatogenesis research (Guo et al., 2017; Green et al., 2018; Wang et al., 2018; Lau et al., 2020; Shami et al., 2020). Thus, similar to the approach used for in vitro oogenesis, these unbiased searches for critical molecules are expected to help identify key factors that facilitate meiosis in vitro (Makar and Sasaki, 2020; Hamazaki et al., 2021).

In contrast, scRNA-seq may not be very effective for investigating post-meiotic stages (also called spermiogenesis), especially mid-spermiogenesis when chromatin begins to condense, because the general transcription machinery is shutting down in this stage (Boussouar et al., 2014). Furthermore, the unique structure of chromatin and its dynamic changes make it difficult to apply common experimental techniques to spermatids. In particular, the highly packed and locus-dependent heterogeneous chromatin state in spermatozoa (i.e., mature sperms) prevents proper biochemical solubilization (Carone et al., 2014; Yamaguchi et al., 2018). Since the failure of chromatin condensation leads to disorganization of not only chromatin but also the entire nucleus and is directly linked to infertility, a better understanding of the sperm chromatin condensation process is essential for both basic and applied research.

NUCLEAR FACTORS FOR HISTONE–PROTAMINE REPLACEMENT

Genome-wide chromatin condensation, which occurs during spermiogenesis, is an epigenetic event unique to sperm that is not observed in other types of cells. Although this is the final step in spermatogenesis, it is likely initiated during meiosis and accompanied by the incorporation and removal of sperm-specific histone variants and histone modifications. After these preparatory steps, chromatin eventually condenses to a higher order as most of the histones in the chromatin are replaced by protamines. This process, i.e., the internal regulation of chromatin for histone–protamine replacement, will be outlined from three major perspectives, “Control of protamine expression and incorporation”, “Histone acetylation for histone–protamine replacement” and “Histone acetylation and DNA repair for histone degradation”.

Control of protamine expression and incorporation

The most critical event for sperm chromatin condensation is the replacement of histones with protamines during mid-spermiogenesis. Protamines are small, basic proteins that are exclusively and abundantly expressed in spermiogenic germ cells. In certain mammals, three protamine genes, Protamine 1 (PRM1/Prm1), Protamine 2 (PRM2/Prm2) and Protamine 3 (PRM3/Prm3), are adjacently situated as a cluster, for example in Chr. 16 (human and mouse) and Chr. 25 (cow). PRM1 protein is synthesized as a mature form, whereas PRM2 is generated by the proteolysis of the PRM2 precursor. Both PRM1 and PRM2 are highly enriched for cysteine and arginine, the former of which generates intra- and inter-protamine disulfide bonds and zinc bridges, resulting in a highly compacted nucleoprotamine complex. In contrast, PRM3 is not enriched in cysteine or arginine, but is characterized by the presence of multiple glutamates at the C-terminus, and Prm3 deletion in mouse has no effect on fertility (Grzmil et al., 2008). All mammalian species possess PRM1 in sperms, while PRM2 is expressed only in a few mammalian species such as humans and rodents (Miller et al., 2010). PRM1 alone is sufficient to condense somatic cell chromatin when it is overexpressed (Iuso et al., 2015), and heterozygous deletion of the Prm1 gene in mice causes male infertility, suggesting that the gene is haploinsufficient (Cho et al., 2001; Mashiko et al., 2013). In contrast, homozygous but not heterozygous deletion of Prm2, by creating nonsense mutations in exon 1 by CRISPR-Cas9, results in impaired chromatin condensation, acrosome formation and motility in mouse sperms and causes infertility (Schneider et al., 2016). Importantly, PRM1 was successfully expressed and incorporated into chromatin in Prm2 homozygous knockout mice (Schneider et al., 2016). Considering the reported PRM1:PRM2 ratio of approximately 2.3:1 in mouse sperms (Balhorn et al., 1977), it seems important to maintain a minimal essential amount of PRMs to sustain fertility.

In humans and mice, PRM1/Prm1–PRM2/Prm2–PRM3/Prm3 genes exist as a cluster. In the promoters of these genes, motif sequences for certain transcription factors such as the TATA-box, cAMP response element (CRE), A-box, B-box and E-like box are located at similar positions and conserved between species (Wykes and Krawetz, 2003). In humans, the –190C→A polymorphism located in the E-α box in the PRM1 promoter is known as a common mutation causing abnormal sperm morphology and an increased PRM1:PRM2 ratio (Gázquez et al., 2008), as well as idiopathic oligozoospermia (Jamali et al., 2016). The CRE motif is targeted by cAMP response element modulator (CREM), a general transcriptional activator that is highly expressed in postmeiotic cells. In Crem knockout mice, spermatogenesis stops at the first postmeiotic stage concomitant with an increase of apoptotic germ cells (Blendy et al., 1996; Nantel et al., 1996). The transcriptional activity of CREM is also regulated by ACT (activator of CREM in the testis) (Steger et al., 2004). Other spermatid-specific small basic proteins are Transition protein 1 and 2 (TNP1 and TNP2). Although the TNP2/Tnp2 gene is situated adjacent to the PRM1/Prm1–PRM2/Prm2–PRM3/Prm3 cluster, while the TNP1/Tnp1 gene is located separately, the promoters of both genes contain similar motif sequences for general transcription factors, such as a TATA-box and a CRE, at similar positions (Wykes and Krawetz, 2003; Miyagawa et al., 2005). Interestingly, KDM3A/JHMD1A/JHDM2A, a histone H3K9 demethylase, targets the promoters of Prm1 and Tnp1, and activates the expression of these genes through histone demethylation (Okada et al., 2007). In addition, KDM3A/JHMD1A/JHDM2A ensures Act expression, and further helps the recruitment of CREM to the Tnp1 and probably also the Prm1, Prm2 and Tnp2 promoters (Liu et al., 2010b), suggesting that epigenetic regulation plays a crucial role in spermiogenesis by controlling the expression of key spermiogenic factors.

Histone acetylation for histone–protamine replacement

Although overexpression of exogenous PRM1 in somatic cells is sufficient to condense their chromatin (Iuso et al., 2015), it is not, alone, able to recapitulate highly ordered sperm histone–protamine replacement. Several events including delayed translation and post-translational modifications of PRMs contribute to highly ordered sperm histone–protamine replacement, and the epigenetic changes on histones for their upcoming removal (and selective retention) are also essential (Fajardo et al., 1997; Wu et al., 2000; Itoh et al., 2019). Recent proteomics analyses identified more than 100 post-translational modifications on histones including methylation, acetylation, phosphorylation, ubiquitylation and oxidation (Luense et al., 2016). Among these modifications, histone acetylation has been studied the most in the context of histone–protamine replacement.

Hyperacetylation of the N-terminal tail of histone H4 is an event that coincides with the time when the round spermatid chromatin begins condensing. The CBP/p300 acetyltransferases are involved in many biological processes and were suggested as being critical for sperm chromatin condensation due to their abundant expression in elongating spermatids (Goodman and Smolik, 2000; Bedford et al., 2010). However, partial depletion of CBP/p300 in elongating spermatids did not affect histone acetylation (Boussouar et al., 2014), although it remains possible that the residual CBP/p300 was sufficient to maintain the acetylation level. A recent study demonstrated that NUT, a nuclear protein in testis, recruits CBP/p300 to enhance H4K5/K8 hyperacetylation, which is required for histone–protamine replacement (Shiota et al., 2018). In addition, genetic deletion of Epc1 or Tip60, components of the NuA4–TIP60 complex, in mice resulted in impaired histone H4 hyperacetylation and in spermiogenic arrest at the round spermatid stage (Dong et al., 2017) (Fig. 1). Intriguingly, both EPC1 and TIP60 are exclusively localized at the atypical pole under the acrosome, showing a cap-like structure, suggesting the spatial regulation of H4 acetylation (Dong et al., 2017). Similar subnuclear localization was also reported for the spermatid-specific histone H1 variant H1T2 and the BET family protein BRD4 (Fig. 1). H1t2 mutant mice showed delayed nuclear condensation and aberrant elongation due to the reduced incorporation of PRMs, and subsequently the presence of residual cytoplasm, acrosome detachment, and fragmented DNA (Martianov et al., 2005; Tanaka et al., 2005; Catena et al., 2006). In contrast, Brd4 knockout causes embryonic lethality, and germ cell-specific knockout has not been reported (Houzelstein et al., 2002). Unlike EPC1, which is not actively involved in gene transcription, BRD4 also functions as a transcriptional activator (Bryant et al., 2015; Dong et al., 2017). ChIP-seq analyses demonstrated that BRD4 and BRDT, a testis-specific BRD family protein (Pivot-Pajot et al., 2003), are enriched in gene promoters in round spermatids (Bryant et al., 2015). Notably, BRD4 and BRDT tend to bind to different subsets of active spermatogenic promoters, unlike CBP/p300, which possess a preference for metabolic promoters (Boussouar et al., 2014). Although H4K16ac (acetylation of Lys16 of histone 4) is not enriched in active promoters, BRD4 is physically bound to H4K5/8/12/16ac, but these acetylated histones subsequently dissociate from BRD4 during sperm elongation and become localized adjacent to the BRD4 ring (Bryant et al., 2015) (Fig. 1), and finally enriched in the central region of spermatozoa, where (peri-)centromeric DNAs also accumulate (Govin et al., 2007).

Fig. 1.

Schematic structure of an elongating spermatid and subnuclear gradation of acetylated histone H4. (A) Exterior appearance (left) and cross section (right) of an elongating spermatid. The spermatid nucleus is surrounded by the acrosome and manchette. NPCs are mainly localized on the distal side. LBR is transiently associated with P1 (Mylonis et al., 2004). In the cross section, the relative amount of acetylated histone H4 (H4ac) is indicated by a color gradient. (B) Enlarged cross-sectional view of the apical region of an elongating spermatid. EPC1/TIP60, BRD4 and H1T2 are localized on the apical side of the nucleus underneath the acrosome (Martianov et al., 2005; Tanaka et al., 2005; Bryant et al., 2015; Dong et al., 2017). EPC1/TIP60 acetylates histone H4 (drawn as blue semicircles with orange stars) before it forms nucleosome structures, and nucleosomes containing hyperacetylated H4 subsequently move to the distal side of the nucleus (Dong et al., 2017). BRD4 also participates in H4 hyperacetylation. BRD4 translocates to both chromatin for transcription and the acrosomal ring (Bryant et al., 2015). NPC, nuclear pore complex; LBR, lamin B receptor; P1, protamine 1; CM, cytoplasmic membrane; OAM, outer acrosome membrane; IAM, inner acrosome membrane, NE, nuclear envelope.

In addition to acetylation, diverse acylations such as propionylation, butylation and crotonylation are reported in male germ cells (Chen et al., 2007; Luense et al., 2016). In spermatids, it has been demonstrated that acetylation and butylation compete for H4K5 and H4K8 (Goudarzi et al., 2016). Since butylation inhibits BRDT binding, this suggests that the usage of acetylation and butylation defines the regions of histone retention and removal. Other acylations are, like acetylation, also localized in transcriptionally active regions. For instance, in spermatids, crotonylation accumulates at transcriptional start sites of sex-linked genes that are activated in an RNF8-dependent manner, and a chromatin conformational change is associated with RNF8-dependent epigenetic programming (Sin et al., 2012). However, RNF8 is not required for H4K16ac or PRM incorporation in spermatids, indicating the distinct functions of acetylation and crotonylation (Abe et al., 2021). Indeed, the sensitivity of these acylations to deacetylases such as HDAC and SIRT1/2 is different from that of acetylation, and they probably function more prominently in situations where acetylation does not function properly (Rousseaux and Khochbin, 2015).

Although histone acetylation in spermatids is likely associated with gene transcription, similar to somatic cells, the unique subnuclear localization of acetyltransferases and acetylated histones suggests that histone–protamine replacement has a certain spatial regularity that is essential for proper chromatin condensation. Several studies have investigated chromosome architecture in sperms by fluorescence in situ hybridization (FISH). These studies proposed that the two chromosome arms juxtapose in an anti-parallel manner, and each chromosome has a hairpin structure on a center–periphery axis (Mudrak et al., 2005). Interestingly, the position of each chromosome appears to be specific along with the accumulation of centromeres in the central region of the sperm nucleus, and their relative localization is preserved among sperm cells of an individual (Millan et al., 2012; Mudrak et al., 2012). It should be noted that application of FISH to the sperm nucleus requires artificial chromatin decondensation to allow the probes to access their target sequences, and this procedure may disrupt the original chromosome architecture to some extent. Despite this caveat, it is likely that sperm chromosome organization is precisely controlled. Furthermore, the sperm chromosome organization appears to be established during the meiotic stages of spermatogenesis (Mudrak et al., 2012). This finding may have some molecular similarities with the notion that the impaired deposition of modified histones during meiosis persists in the post-meiotic stages (Maezawa et al., 2018).

Impaired sperm chromosome organization has been reported in infertile men (Finch et al., 2008). Although it is probably impractical to apply FISH for routine sperm examinations in assisted reproduction technology, precise measurement of nuclear factors to assess sperm quality has been in increasing demand in recent years. For instance, in human sperms, large nuclear vacuoles are sometimes observed in the apical side of the sperm head, which coincide with reduced DNA condensation (Boitrelle et al., 2011) (Fig. 2), lower pregnancy rates, and higher abortion rate (Berkovitz et al., 2006). However, other studies have shown that sperm nuclear vacuoles are a normal physiological feature and do not affect fertility (Fortunato et al., 2016; Morin and Scott, 2018). Perhaps the reason for this discrepancy is that at the laboratory level, it is difficult to accurately distinguish between sperm nuclear vacuoles and sperm surface indentations and measure the size of the nuclear vacuoles. Thus, collecting more cases based on accurate metrics is essential to determine the effect of sperm nuclear vacuoles on male fertility.

Fig. 2.

Schematic illustration of an elongating spermatid with impaired nuclear structure. (A) Normal (left) and three types of sperm nuclear malformations: large nuclear vacuolation (middle left), acrosome fragmentation/hypoplasia (middle right) and manchette elongation (right). Abnormal acrosome biogenesis mainly causes nuclear deformation and occasionally affects chromatin condensation. Abnormal manchette elongation causes a knob-like nuclear structure, in which chromatin condensation is sometimes impaired on the distal side. (B) Immunostaining and confocal microscopic view of a human sperm containing a nuclear vacuole (arrow). The vacuole exhibits low DNA (left) and accumulation of histone H3 (green, middle). PRM2 (red, right) is present to some extent. The immunostaining images were kindly provided by Dr. Satoshi Kaneko from Ichikawa General Hospital, Tokyo Dental College.

Histone acetylation and DNA repair for histone degradation

Notably, histone acetylation during spermiogenesis is also critical for the massive degradation of histones removed from the chromatin. In general, H4K5/8/12ac are highly associated with transcriptional activation, whereas H4K16ac is strongly linked to open chromatin regions as it blocks H4 tail binding to the acidic patch of the nucleosome and prevents compact chromatin formation (Shogren-Knaak et al., 2006; Robinson et al., 2008; Potoyan and Papoian, 2012; Kalashnikova et al., 2013). Consistent with these observations, in equine spermatogenesis, only H4K16ac accumulates in elongating spermatids, whereas H4K5/8/12ac are seen in the earlier stages, suggesting a specific effect of H4K16ac on histone–protamine replacement (Ketchum et al., 2018). In mice, H4 hyperacetylation and DNA strand breakage occur almost simultaneously and transiently in mid-spermiogenesis (H4ac in Step 9–12, TUNEL staining in Step 9–11, and γH2A.X in Step 9–12) (Lahn et al., 2002; Marcon and Boissonneault, 2004; Jha et al., 2017). Approximately 5–10 million lesions are expected to be generated genome-wide in mid-spermiogenesis, and this step-specific DNA strand breakage, generated by Topoisomerase II beta (TOP2B), is assumed to be intrinsic, allowing a change in DNA topology, although many other extrinsic factors such as reactive oxygen species (ROS) can cause DNA damage (Chen and Longo, 1996; Laberge and Boissonneault, 2005). Nucleosome withdrawal from DNA presumably leaves several unconstrained negative DNA supercoils in the topological domain and they must be eliminated by the introduction of single- or double-strand breaks in DNA (Boissonneault, 2002). In the late spermiogenic stages and thereafter, including mature sperm, DNA strand breaks are no longer detected, however, and at this time poly(ADP-ribose) polymerase 1 (PARP1) and poly(ADP-ribose) glycohydrolase (PARG) play crucial roles. PARP1 binds TOP2B as it cleaves DNA, and then undergoes poly(ADP-ribosyl) ation (Meyer-Ficca et al., 2011). This causes TOP2B to release the DNA while completing its DNA strand passage. PARG then digests the poly(ADP-ribose) from PARP1, which, in turn, separates PARP1 from TOP2B, allowing the cycle to repeat itself (Meyer-Ficca et al., 2011; Ward, 2011). These observations imply that such genome-wide DNA strand breaks are properly repaired along with the DNA condensation and histone–protamine replacement, even though inefficient non-homologous end joining (NHEJ) is the only repair pathway available during spermiogenesis (Ahmed et al., 2015). The functional existence of the PARP1–XRCC1-dependent NHEJ pathway was successfully demonstrated in round spermatids (Ahmed et al., 2010), while PARP1 inhibition is dispensable for the repair of irradiation-induced DNA strand breaks in elongated spermatids (Ahmed et al., 2015). TNP1 and TNP2 were also expected to be involved due to their DNA repair and ligation stimulation activities on short DNA fragments in vitro (Lévesque et al., 1998; Caron et al., 2001). The contribution of TNPs to DNA repair has not been examined further in vivo, although it was recently demonstrated that TNPs do not directly displace histones from chromatin, but rather mediate the recruitment and processing of PRMs through binding to H2A.L.2 (Barral et al., 2017).

Evicted histones are subjected to proteasome degradation, and PA200, a proteasome activator that is preferentially expressed in testis, plays a critical role in degrading histones (Qian et al., 2013). Deletion of PA200 in mice results in a delay of core histone removal in elongated spermatids. Importantly, purified PA200 promotes proteasomal degradation of acetylated core histones in response to DNA strand breaks, but not of polyubiquitinated proteins, further supporting the critical role of histone acetylation rather than ubiquitination for sperm histone degradation, although H3 ubiquitination by PHF7 indirectly contributes to histone removal by BRDT (Kim et al., 2020).

NON-NUCLEAR FACTORS INVOLVED IN SPERM NUCLEAR CONDENSATION

In many knockout mice studies, impaired chromatin condensation is frequently associated with abnormal sperm head morphology. Failure of chromatin condensation generally induces round-headed sperm in mice, but what if the impairment of sperm head formation occurs first? During mid-spermiogenesis, various extranuclear structures are specifically formed so that nuclear remodeling may proceed (Pereira et al., 2019; Teves et al., 2020). The anterior side of the spermatid head is covered by an acrosome, while the posterior side is surrounded by a manchette. Perinuclear structural transformation is associated with changes in the composition and organization of the nuclear lamina and redistribution and elimination of nuclear pore complexes (NPCs), and, further, chromatin condensation. Although a direct causal relationship between perinuclear structural transformation and chromatin condensation is not always demonstrated, it is an interesting possibility that chromatin condensation is regulated by extrinsic factors such as nuclear forces, perinuclear structure, and possibly nuclear volume. Three non-chromatin structures – the acrosome, the manchette, and NPCs/nucleoporins/nuclear envelope – are substantially involved in this process, and the essence of each factor is described below.

Acrosome

The acrosome is an organelle with a cap-like structure that develops over the anterior half of the head of the sperm in many animals. It is derived from the Golgi apparatus and contains digestive enzymes that are required for penetration of the zona pellucida during fertilization. Thus, acrosomal defects directly cause male infertility due to the failure of fertilization, notably in a specific clinical condition called globozoospermia, which is characterized by round-headed sperm with impaired acrosome formation. By September 2021, seven genes, DPY19L2, SPATA16, PICK1, CSNK2A2, GOPC, SPACA1 and GBA2, had been listed as genetic factors causing globozoospermia in the OMIM (Online Mendelian Inheritance in Man) database (https://omim.org/). Among these genes, mutations in DPY19L2 account for more than 70% of cases (Liu et al., 2010a; Harbuz et al., 2011; Koscinski et al., 2011; ElInati et al., 2016; Ghédir et al., 2016; Ray et al., 2017). Knockout mice have been established for all of these genes, and male infertility phenotypes have been reported (Xu et al., 1999; Suzuki-Toyota et al., 2004; Yildiz et al., 2006; Xiao et al., 2009; Fujihara et al., 2012, 2017; Pierre et al., 2012). In the Dpy19l2 knockout mice, the nuclear shape of spermatids is severely distorted and causes infertility, whereas the influence on chromatin condensation seems to vary among the models (Fig. 2). Impaired chromatin condensation has been occasionally observed in Dpy19l2 knockout mice (Pierre et al., 2012), but it also varied among the individual spermatids/sperms, likely depending on the severity of nuclear deformation. Disruption of Pick1 in mice induces male infertility, and the phenotype resembles human globozoospermia (Xiao et al., 2009). In addition to the fragmentation of acrosomes, impaired nuclear elongation and chromatin condensation are also reported in the maturation phase of spermatids in Pick1 knockout mice (Xiao et al., 2009). In contrast, chromatin condensation appears normal in Gopc (Golgi-associated PDZ- and coiled-coil motif-containing protein) and Mfsd14a (an MFS transporter) knockout mice (Suzuki-Toyota et al., 2004; Doran et al., 2016), even though GOPC interacts with PICK1 (Xiao et al., 2009). Furthermore, mice carrying a patient-type point mutation in Spata16 maintain their fertility, while exon deletion of Spata16 resulted in infertile male mice due to spermiogenic arrest rather than a globozoospermia-like phenotype (Fujihara et al., 2017), suggesting that proper acrosome formation is linked to, but not always essential for, sperm chromatin condensation (Fig. 2).

Recently, a member of the histone deacetylases, SIRTUIN, was revealed to regulate acrosome biogenesis. Germ cell-specific Sirt1 knockout mice were infertile and initially reported to exhibit low sperm count and an increased proportion of abnormal spermatozoa (Bell et al., 2014). Despite the function of SIRT1 as a histone deacetylase, however, H4 acetylation was significantly lower in the knockout testis, implying an indirect effect of SIRT1 in altering histone acetylation (Bell et al., 2014). Later, the spermiogenic phenotype of Sirt1 knockout mice was reported to resemble human globozoospermia due to defective acrosome formation (Liu et al., 2017). It is notable that the deacetylation of autophagy factors LC3 and ATG7 was disrupted in Sirt1 knockout spermatids, causing the failure of LC3 to be recruited to Golgi apparatus-derived vesicles as well as the failure of GOPC and PICK1 to be recruited to acrosomal vesicles (Liu et al., 2017). Similarly, Sirt6 knockout mice displayed arrested spermatid elongation with impaired acrosome formation, whereas the deacetylation activity on histone H3 was found to be dispensable for spermatogenesis (Wei et al., 2020), suggesting that SIRTUIN has a distinct, non-chromatin function in acrosome biogenesis.

Manchette

During spermatid elongation, protein transport is required for proper formation of the sperm head, and a microtubule-based protein delivery platform, the manchette, provides essential support for this process. The manchette is a skirt-like cytoskeletal structure consisting of α-tubulin and several trafficking proteins (Soley, 1997; Lehti and Sironen, 2016). It is specifically formed in the posterior half of the sperm head during spermatid elongation, and when the manchette is absent, the nuclear shape becomes abnormally round. The manchette is physically connected to the nucleus through the LINC (linker of nucleoskeleton and cytoskeleton) complex, which includes SUN and KASH/Nesprin (Bouzid et al., 2019; Manfrevola et al., 2021). In other words, for manchette formation, the LINC complex functions in coupling nuclear structures and the surrounding cytoskeleton (Crisp et al., 2006).

SUN3, SUN4 and SUN5 have been demonstrated to play crucial roles in spermatid elongation through knockout studies (Pasch et al., 2015; Gao et al., 2020; Zhang et al., 2021). SUN3 and SUN4 proteins are completely colocalized in the manchette and bind to each other. Mice with either gene deleted are both infertile and show a globozoospermia-like phenotype, owing to the failure of spermatid elongation, pointing to a cooperative role for SUN3 and SUN4 in transferring cytoplasmic force to shape the sperm nucleus. However, chromatin condensation is not affected in either knockout mouse (Pasch et al., 2015; Gao et al., 2020). On the other hand, SUN5 possesses a distinct function from SUN3/4: its knockout causes detachment of the sperm head from the tail due to the absence of basal plates (Zhang et al., 2021). This observation is consistent with the phenotype observed in human patients carrying homozygous deletion of SUN5 (Elkhatib et al., 2017b). Also in the Sun5 knockout mice, sperm chromatin condensation appears normal, at least in transmission electron microscopic analyses (Zhang et al., 2021).

Microtubule tracks and motor proteins including actins and kinesins are also major components of the manchette. Thus, many mouse models bearing mutations in microtubule-related proteins exhibit deformed nuclei in their elongated spermatids (Lehti et al., 2013; Lehti and Sironen, 2016; Teves et al., 2020). Unlike the defects of LINC proteins, the zipper-like movement of the manchette is mainly affected in these cases, and the nucleus tends to form a knob-like shape with an abnormally elongated manchette rather than a round shape (Lehti and Sironen, 2016) (Fig. 2). In knockout mice of Kif3a or Lis1, with Lis1 reported to bind the dynein complex, defective chromatin condensation is apparent in the posterior region where an abnormally elongated manchette is formed (Nayernia et al., 2003; Lehti et al., 2013), implying an effect of nuclear volume on chromatin condensation (Fig. 2).

NPCs/nucleoporins/nuclear envelope

Nuclear pore complexes are conventionally known to regulate nucleocytoplasmic transport. Structurally, the NPC consists of ~30 discrete Nucleoporins (Nups) constitutively organized to create cytoplasmic, inner and nuclear regions within the NPC (Beck and Hurt, 2017). Recent studies have demonstrated that NPCs interact with the genome and contribute to regulating gene expression (Kuhn and Capelson, 2019; Pascual-Garcia and Capelson, 2021). NPCs are involved in both transcriptional activation and repression depending on their associating chromatin factors. Dynamics of the NPCs mainly play a role in later stages of spermatid maturation, based on the observation of a global redistribution of NPCs to the redundant nuclear envelope compartment in developing spermatids in mice (Ho, 2010). During nuclear condensation of spermatids, distribution patterns of nuclear pores are greatly affected by the developing acrosome and manchette (Ho, 2010).

Among the numerous components of NPCs, some have been reported to exhibit spermatogenic defects in knockout mice. RanBP1 is a cofactor of the small GTPase Ran, and plays crucial roles in nucleocytoplasmic transport and nuclear envelope formation (Yoneda, 2000; Hetzer et al., 2002; Weis, 2003). Ranbp1 knockout mice exhibit male infertility owing to a mid-spermiogenic defect, whereas the females maintain fertility (Nagai et al., 2011). Ablation of KNAP4/Importin α4, but not KNAP3/Importin α3, causes male subfertility with abnormal sperm head morphology concomitant with altered expression of spermatogenesis-related proteins, suggesting its role in maintaining chromatin state and gene expression in spermiogenesis (Miyamoto et al., 2020). Furthermore, whole-exome sequencing revealed a homozygous mutation in the testis-specific gene Nucleoporin 210 like (NUP210L) in an infertile patient with large sperm heads owing to highly uncondensed chromatin (Arafah et al., 2021). ELYS is also a well-studied NPC component. It was originally identified as a transcription factor because of its ability to induce gene expression (Kimura et al., 2002) and is the only Nup possessing nucleosome-binding capacity (Inoue and Zhang, 2014; Zierhut et al., 2014). However, the function of ELYS in spermiogenesis has not been tested due to the embryonic lethality of knockout mice.

Considering the diverse functions of the nuclear envelope in both nuclear shape and chromatin dynamics (Karoutas and Akhtar, 2021), it is not surprising that the components of the nuclear envelope are also highly involved in sperm nuclear condensation (Pereira et al., 2019). In particular, lamin B receptor (LBR) temporally associates with PRM1 in mouse elongating spermatids depending on the phosphorylation of PRM1, illustrating a model of sequential binding of PRM1 to LBR and then to DNA during sperm chromatin condensation (Mylonis et al., 2004) (Fig. 1). However, LBR is not expressed in human spermatids, and LEMD1, LAP2β and lamin B1 (LMNB1), and their temporal expression during spermiogenesis, have been proposed to substitute for LBR in human sperm although further investigations are essential (Elkhatib et al., 2015, 2017a). Unlike LBR, which localizes to the posterior pole of elongated spermatids (Mylonis et al., 2004), localization of lamin A/C (LMNA/C) shifts from the nuclear membrane to the acroplaxome, a structure found between the acrosomal membrane and the nuclear membrane, during spermiogenesis. Reduction of LMNA level by testicular siRNA injection leads to a deficiency in assembling the acroplaxome and disturbs the formation of the manchette, resulting in a deformed sperm nuclear shape (Shen et al., 2014). This implies a functional connection between nuclear envelope, acrosome and manchette.

As mentioned above, it is apparent that these three non-chromatin components – acrosome, manchette and NPCs/Nups/nuclear envelope – are indirectly and cooperatively involved in sperm nuclear condensation. Thus, it would be interesting to reconstitute nucleoprotamine (i.e., the DNA–PRM complex) in an artificial closed space of various sizes that mimics the reduction in nuclear volume during spermiogenesis (Fig. 3). The attempt to encapsulate DNA–PRM complexes in nanoparticles has been studied for many years as a method for the artificial delivery of nucleic acids into cells, including mRNA vaccines (Ruseska et al., 2021). This was originally inspired by spermatogenesis in nature, but recent developments in nanoparticle research may now be applied to spermatogenesis research in the opposite direction.

Fig. 3.

Does nuclear volume control sperm nuclear condensation? (A) In an elongating spermatid, the nucleus is surrounded by the acrosome from the apical side, and the manchette and nuclear envelope from the basal side, reducing its volume accompanied by histone–protamine replacement. (B) Application of nanoparticles for in vitro reconstitution of histone–protamine replacement. Certain types of nanoparticles can change their volume as desired. Thus, utilizing these particles may mimic nuclear compaction during spermiogenesis and allow us to examine the effect of nuclear size on chromatin condensation.

APPLICATIONS OF NGS AND SINGLE-CELL ANALYSES TO SPERMATOLOGY

In the past decade, high-throughput analyses such as next-generation sequencing (NGS) have remarkably contributed to the understanding of sperm chromatin structure at the genome-wide level. For sperm chromatin, MNase-seq and ChIP-seq have been frequently applied to identify the localization of retained nucleosomes, and these studies were able to demonstrate the modification-dependent retention of histones in specific genomic elements (Saitou and Kurimoto, 2014; Okada and Yamaguchi, 2017; Yamaguchi et al., 2018). More recently, NGS started to be utilized for investigating higher-order chromatin structure of sperms, such as in the Hi-C method, as the folding principle of the sperm genome is poorly understood. Unexpectedly, however, the high-order chromatin structure of sperms suggested by Hi-C was essentially similar to that of somatic cells and ES cells, except for more frequent proximity to distant locations on the same chromosome and between different chromosomes (Battulin et al., 2015; Jung et al., 2017; Ke et al., 2017; Wang et al., 2019). Although these results are consistent with the characteristics of highly condensed sperm chromatin, it is somewhat difficult to imagine that the sperm genome has almost the same higher-order structure as the somatic genome, in a nuclear volume only several tenths that of a somatic cell (Ward and Coffey, 1991), and with the loss of more than 90% of histones (Erkek et al., 2013). Another study involving Hi-C demonstrated that the sperm genome does not form distinct topologically associating domains like somatic cells, although compartmentalization associated with active transcription is maintained (Vara et al., 2019). The cause of these discrepancies is unclear; however, it is possible that experimental procedures were not adequately adapted for sperms. Further comprehensive analyses including more than just sequencing are necessary for precise verification.

Nevertheless, although some technical improvements seem to be required, NGS is undoubtedly a powerful tool for sperm chromatin study. In particular, the application of single-cell analysis tools will be even more important in spermatology than in research on other cell types, considering that a single individual is born from a single sperm and the remarkable heterogeneity of sperms in non-laboratory animals including humans. In a recent demonstration of the power of this approach, single-cell RNA-seq combined with precise bioinformatic analyses succeeded in elucidating sperm-level natural selection and evolutionary conflict (Bhutani et al., 2021). One of the next important research questions is how to apply the findings of these basic studies on sperm chromatin to clinical sperm testing. In this regard, appropriate translational research is essential and will be beneficial for the upcoming insurance coverage of assisted reproduction technology in Japan.

ACKNOWLEDGMENTS

I would like to thank all members of the Laboratory of Pathology and Development from the Institute for Quantitative Biosciences for insightful discussions. I also thank Dr. Satoshi Kaneko from Ichikawa General Hospital for providing the immunostaining images shown in Fig. 2, and Dr. Satoko Arakawa from Tokyo Medical and Dental University for sharing her expertise. This work was supported by JSPS KAKENHI Grant Numbers 19H05254 and 20H05939, and JST ERATO Grant Number JPMJER1901.

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