TY - JOUR
AU - O’Bryan, Moira K
AB - Abstract BACKGROUND Members of the cysteine-rich secretory proteins (CRISPS), antigen 5 (Ag5) and pathogenesis-related 1 (Pr-1) (CAP) superfamily of proteins are found across the bacterial, fungal, plant and animal kingdoms. Although many CAP superfamily proteins remain poorly characterized, over the past decade evidence has accumulated, which provides insights into the functional roles of these proteins in various processes, including fertilization, immune defence and subversion, pathogen virulence, venom toxicology and cancer biology. OBJECTIVE AND RATIONALE The aim of this article is to summarize the current state of knowledge on CAP superfamily proteins in mammalian fertility, organismal homeostasis and disease pathogenesis. SEARCH METHODS The scientific literature search was undertaken via PubMed database on all articles published prior to November 2019. Search terms were based on following keywords: ‘CAP superfamily’, ‘CRISP’, ‘Cysteine-rich secretory proteins’, ‘Antigen 5’, ‘Pathogenesis-related 1’, ‘male fertility’, ‘CAP and CTL domain containing’, ‘CRISPLD1’, ‘CRISPLD2’, ‘bacterial SCP’, ‘ion channel regulator’, ‘CatSper’, ‘PI15’, ‘PI16’, ‘CLEC’, ‘PRY proteins’, ‘ASP proteins’, ‘spermatogenesis’, ‘epididymal maturation’, ‘capacitation’ and ‘snake CRISP’. In addition to that, reference lists of primary and review article were reviewed for additional relevant publications. OUTCOMES In this review, we discuss the breadth of knowledge on CAP superfamily proteins with regards to their protein structure, biological functions and emerging significance in reproduction, health and disease. We discuss the evolution of CAP superfamily proteins from their otherwise unembellished prokaryotic predecessors into the multi-domain and neofunctionalized members found in eukaryotic organisms today. At least in part because of the rapid evolution of these proteins, many inconsistencies in nomenclature exist within the literature. As such, and in part through the use of a maximum likelihood phylogenetic analysis of the vertebrate CRISP subfamily, we have attempted to clarify this confusion, thus allowing for a comparison of orthologous protein function between species. This framework also allows the prediction of functional relevance between species based on sequence and structural conservation. WIDER IMPLICATIONS This review generates a picture of critical roles for CAP proteins in ion channel regulation, sterol and lipid binding and protease inhibition, and as ligands involved in the induction of multiple cellular processes. Male fertility, male infertility, sperm, sperm function, pathological disorder, reproduction, male reproductive tract, epididymis, gamete biology, sperm maturation Introduction The cysteine-rich secretory proteins, antigen 5, and pathogenesis-related 1 (CAP) superfamily proteins (pfam PF00188) are found in diverse species across all kingdoms (Fry et al., 2009; Abraham and Chandler, 2017). The three eponymous subfamilies—the cysteine-rich secretory proteins (CRISPs) in vertebrates, antigen 5 (Ag5) related proteins in insects and pathogenesis-related 1 (Pr-1) proteins in plants, and all other members of the CAP superfamily, share significant sequence homology defined by the possession of a conserved CAP domain (Gibbs et al., 2008). The majority of CAPs also possess a hinge region linked to one of multiple alternative C-terminal domains. While the function of virtually all CAP proteins remains poorly understood, they are known to play significant roles in processes including sperm development and function, immune defence in mammals and plants and pathogen virulence; as venoms; and in cancer biology. Each of their roles will be reviewed herein. Genomic comparisons reveal that there are 49 CAPs in humans and 33 in mice (InterPro Protein Domain: IPR035940). Comparative analyses also indicate that the CAP superfamily can be subdivided in 11 subfamilies: CRISPs, glioma pathogenesis-related (GLIPR) proteins, the Golgi-associated pathogenesis-related (GAPR) proteins, the cysteine-rich LCCL domain-containing (CRISPLD) proteins, the peptidase inhibitor 15 (PI15) proteins, the peptidase inhibitor 16 (PI16) proteins, the C-type lectin (CLEC) proteins, the venom allergen proteins (Ag), the bacterial SCP domain-containing proteins, the pathogenesis-related (Pr) proteins and the fungal pathogenesis-related (PRY) proteins (Abraham and Chandler, 2017) (Fig. 1). Due to an incomplete view of the breadth of the superfamily at the time of individual sequence discovery, however, some members are often referred to by inaccurate nomenclature, thus causing confusion within the literature. For example, PI15, PI16 and CRISPLD1 proteins have been referred to as CRISP8, CRISP9 and CRISP10 in public databases although, as explained below, they clearly do not belong to the CRISP subfamily. Figure 1 Open in new tabDownload slide Schematic representation of CAP subfamily protein structure. The schematic representation focuses on the structural composition of 11 subfamilies of the cysteine-rich secretory proteins, antigen 5, and pathogenesis-related 1 (CAP) superfamily. The conserved CAP motifs are highlighted in red (CAP1), purple (CAP2), green (CAP3) and blue (CAP4) in the CAP domain (highlighted in light blue). The hinge domain is highlighted in salmon followed by the subfamily specific C-terminal extension domains. Figure 1 Open in new tabDownload slide Schematic representation of CAP subfamily protein structure. The schematic representation focuses on the structural composition of 11 subfamilies of the cysteine-rich secretory proteins, antigen 5, and pathogenesis-related 1 (CAP) superfamily. The conserved CAP motifs are highlighted in red (CAP1), purple (CAP2), green (CAP3) and blue (CAP4) in the CAP domain (highlighted in light blue). The hinge domain is highlighted in salmon followed by the subfamily specific C-terminal extension domains. In this article, we will review the current state of knowledge on CAP superfamily proteins. We will discuss the evolution of the CAP superfamily and its conserved structural domains. We will undertake an in-depth analysis of the whole CAP superfamily so as to allow insights into the commonality of domain function, and in the hope of unravelling some of the cross-species confusion that currently occurs in the literature. Lastly, we will summarize the role of mammalian CAP superfamily proteins in fertility and their association with a range of pathologies. Evolution of CAP Superfamily Proteins As reflected by the number of distinct CAP subfamilies, considerable sequence variation exists between contemporaneous CAPs. The majority, however, retain a core domain structure, namely a CAP domain linked to a ridged linker domain (hinge), then a variable C-terminal domain that is proposed to have subfamily specific function. The defining feature of the CAP superfamily is the presence of CAP motifs (CAP1–4) in a CAP domain (Gibbs et al., 2008). As detailed below, early evolving CAPs lack the C-terminal domain or both the hinge domain and C-terminal domain. The molecular evolution of the CAP superfamily proteins has recently been analysed (Abraham and Chandler, 2017). Data suggest that the full range of CAP proteins was elucidated via a 14-step process, which initiated in bacteria and is fully elaborated in vertebrates. It is proposed that CAP superfamily proteins first appeared as single CAP domain protein/SCP-like protein with common ancestors in bacteria, as Step 1. Unlike eukaryotic CAP proteins, most of the bacterial CAP proteins lack the CAP2 signature motif and do not contain the usual series of cysteine residues found in the advance CAP domains (Abraham and Chandler, 2017). Functionally, early bacterial-CAP proteins are thought to play a role in modulating host immunity (Hewitson et al., 2009; Asojo et al., 2018). In Steps 2–4, the single domain protein diverged to form the GLIPR2/GAPR subfamily, the Pr subfamily in plants and the fungal PRY subfamily. The majority of the Step 2–4 single-domain CAPs are found in invertebrates, such as round worms and arthropods, where they have been shown to have lipid and sterol-binding capability (Osman et al., 2012; Mohandas et al., 2015; Stroehlein et al., 2016). They differ from Step 1 proteins by the presence of the distinct CAP motifs (CAP1–4) in the CAP domain, as opposed to bacterial CAP proteins which lack the CAP2 motif and four cysteine residues that are conserved across early eukaryotic CAPs (Abraham and Chandler, 2017). In Step 5, the hinge domain, a rigid linker, was added as an N-terminal extension to the CAP domain to form two-domain proteins, which later evolved into the Ag proteins (in Steps 7 and 8). In parallel, and in what the authors refer to a Step 6, additional C-terminal extensions were added to form three-domain proteins—the CRISPs and GLIPR1 subfamilies (Abraham and Chandler, 2017). The third domain (or C-terminal extension) is characteristic for individual CAP subfamilies and is proposed to impart distinct functions (discussed below). Further elaboration of three-domain CAPs leads to the PI15 and PI16 subfamilies, the CRISPLD protein subfamily and the CLEC protein subfamily in vertebrates (Steps 9 and 10). In Steps 11 and 13, the CAPs underwent a major evolutionary expansion as a consequence of gene duplications. For example, the CRISPs expanded to form three to four paralogues in most vertebrates. Notably, CRISP2 further evolved into distinct lineages of CRISP1 and CRISP3 (Step 11) in amphibians, and later in rodents an additional protein, CRISP4 (Step 13) (Abraham and Chandler, 2017). Another example includes the GLIPR genes, in which GLIPR1L3 is the result of a gene duplication event in GLIPR1L1 (Ren et al., 2006). The most recent evolution of CAPs is proposed to have occurred in early mammals with the duplication of the CLEC18A gene (from Step 10) to form CLEC18B (Step 12) and CLEC18C (Step 14) in primates (Abraham and Chandler, 2017). Each CAP subfamily and their function will be discussed in this review. For more details on CAP superfamily evolution and phylogenetic analysis, readers are referred to the literature (Gibbs et al., 2008; Abraham and Chandler, 2017; Vicens, 2018). Of particular relevance to the focus of this review, an extensive body of literature indicates that genes involved in fertilization, and thus speciation, evolve more rapidly than genes involved in body-wide functions (Swanson and Vacquier, 2002; Vicens and Treviño, 2018). This rapid evolution is often driven by positive (Darwinian) selection. While some controversy exists, collectively data indicate that the CRISPs are undergoing positive selection. Specifically, using translated nucleotide sequences, Sunagar et al., (2012) found evidence of positive selection driven evolution of reptilian venom CRISPs, but a negative selection driven evolution of mammalian CRISPs (Sunagar et al., 2012). By contrast, a more comprehensive and domain specific analysis using a larger number of sequences undertaken by Vicens and Treviño revealed that several members of the mammalian CRISP subfamily have evolved under positive selection (as discussed below). Notably there appears to be a stronger influence of positive selection in CRISP3 than in CRISP1 and CRISP2. Specifically, within CRISP1, 18.1% of the CRISP domain residues appear to have undergone positive selection, whereas in CRISP2, the CAP domain was the primarily region of positive selection (Vicens and Treviño, 2018). By contrast, in CRISP3 54.3% of CAP domain residues and 39.4% of the CRISP domain residues were mapped as being under positive selection (Vicens and Treviño, 2018). Collectively, these data suggest, but do not prove, important roles for CRISPs in fertilization and/or speciation. Consistent with this interpretation, there is considerable intra-orthologue sequence variation between species as well as between paralogues. This has been confusing for researchers when attempting to assign an informative name to newly discovered sequences. As a consequence, there is considerable inaccuracy in the naming of orthologous CRISPs between species, and the inappropriate assigning of function based on names rather than orthology. Thus, in this review, for the CRISP proteins, which are the most well characterized of the CAP fertility proteins, we have carried out a phylogenetic analysis to definitively identify orthologues in the hope of being able to more insightfully compare functions at a cell and physiological level. This comparison is found in the relevant sections below. Structural Domains of CAP Superfamily Proteins As indicated above, and confirmed by crystal structures of reptile (stercrisp, natrin, triflin, helothermine) and plant (P14a) CAPs, fully elaborated CAP superfamily proteins typically contain three domains: the CAP domain, the hinge domain and a variable C-terminal domain (Nobile et al., 1996; Fernández et al., 1997; Monsalve et al., 1999; Eberle et al., 2002; Yamazaki et al., 2002; Asojo et al., 2005; Shikamoto et al., 2005; Zhou et al., 2008; Darwiche et al., 2016). In this section, we will briefly discuss the conserved structural features of the CAP domain and, as an exemplar, the hinge and ion channel regulatory (ICR) domains within the CRISP subfamily as a model for all CAP proteins. The structural features of other members of the CAP superfamily are discussed under relevant headings. The CAP domain The defining feature of the CAP superfamily proteins is the presence of a CAP domain containing four highly conserved signature motifs (CAP1–4) (Gibbs et al., 2008). CAP domains are typically 20 kDa in size, approximately 215 amino acids in length, and have variably been called PR-1 (pathogenesis related-1) domains or TAPS (Tpx/antigen 5/pathogenesis-related-1/Sc7) domains (Brooks et al., 1986; van Loon and van Strien, 1999; Osman et al., 2012). The four signature motifs are, CAP1: [GDER][HR][FYWH][TVS][QA][LIVM][LIVMA]Wxx[STN], CAP2: [LIVMFYH][LIVMFY]xC[NQRHS]Yx[PARH]x[GL]N[LIVMFYWDN] (Prosite: http://www.expasy.ch/prosite/), CAP3: [HNxxR] and CAP4: G[EQ]N[ILV] (Figs 2 and 3A) (Gibbs et al., 2008). Together they form a unique structural fold of α–β–α sandwich in which two layers of α-helices flank the central three-stranded anti-parallel β-sheets (Guo et al., 2005; Zhou et al., 2008; Brangulis et al., 2015; Olrichs et al., 2016). These loops are stabilized by internal hydrogen bonding and three crossed disulphide bridges in the bipartite hydrophobic core of the CAP domain (Szyperski et al., 1998; Guo et al., 2005; Wang et al., 2005). The difficulty in accurately predicting the structure of individual CAP domains arises due to high structural variations in non-core loop length and high (47%) α-helix and β-sheet content within each motif, and the formation of disulphide bonds to impart structural stability (Darwiche et al., 2016). Figure 2 Open in new tabDownload slide Sequence alignment of the signature CAP motifs (CAP1–4) across members of the CAP superfamily of proteins. The major conserved secondary structure elements are indicated above the relevant amino acid sequence, with α-helices and β-strands highlighted by black coils and arrows respectively. Conserved amino acids from this alignment are highlighted in a yellow box and individual CAP motifs (CAP1–4) are highlighted within a blue box. The sequence alignment was generated using ENDscript 2.0 and the structural analyses was carried out using PyMOL v2.1.0. Figure 2 Open in new tabDownload slide Sequence alignment of the signature CAP motifs (CAP1–4) across members of the CAP superfamily of proteins. The major conserved secondary structure elements are indicated above the relevant amino acid sequence, with α-helices and β-strands highlighted by black coils and arrows respectively. Conserved amino acids from this alignment are highlighted in a yellow box and individual CAP motifs (CAP1–4) are highlighted within a blue box. The sequence alignment was generated using ENDscript 2.0 and the structural analyses was carried out using PyMOL v2.1.0. Figure 3 Open in new tabDownload slide Schematic of key features of the conserved N-terminal CAP domain of CAP superfamily proteins. A Mapping of the conserved CAP motifs (CAP1–4) on cysteine-rich secretory protein (CRISP) protein (Stecrisp, Protein Data Bank (PDB) ID: 1RC9). The signature motifs are annotated as; CAP1 in red, CAP2 in yellow, CAP3 in green and CAP4 in blue. The proposed CAP cavity is denoted by the purple ball-like structure between the CAP motifs. B Surface plots of the representative members of the CAP superfamily proteins showing conserved CAP cavity. The plots of (a) tomato, P14a (1CFE), (b) natrin (1XX5), (c) stecrisp (1RC9), (d) GAPR-1 (1SMB), (e) Ves v 5 (1QNX), (f) Na-ASP-2 (1U53), (g) Pry1 (5ETE) and (h) pseudechetoxin (2DDA) are shown in the same orientation and the location of the two conserved Glu and His residues are marked red, indicating the CAP cavity. C CAP cavity formed by the conserved amino acid residues across CAP superfamily. The superposed structures of stecrisp (Trimeresurus stejnegeri; 1RC9), red; natrin (Naja atra; 1XX5), cyan; pseudechetoxin (Pseudechis australis; 2DDA) magenta and glioma pathogenesis-related 1 (GLIPR1) (Homo sapiens; 3Q2R), yellow showing key solvent exposed residues corresponding to His60, Glu75, Glu96 and His115 within the representative CAP domain. To superimpose CAP cavity, all the PDB structures were oriented and aligned based on structural similarity using PyMOL v2.1.0. Figure 3 Open in new tabDownload slide Schematic of key features of the conserved N-terminal CAP domain of CAP superfamily proteins. A Mapping of the conserved CAP motifs (CAP1–4) on cysteine-rich secretory protein (CRISP) protein (Stecrisp, Protein Data Bank (PDB) ID: 1RC9). The signature motifs are annotated as; CAP1 in red, CAP2 in yellow, CAP3 in green and CAP4 in blue. The proposed CAP cavity is denoted by the purple ball-like structure between the CAP motifs. B Surface plots of the representative members of the CAP superfamily proteins showing conserved CAP cavity. The plots of (a) tomato, P14a (1CFE), (b) natrin (1XX5), (c) stecrisp (1RC9), (d) GAPR-1 (1SMB), (e) Ves v 5 (1QNX), (f) Na-ASP-2 (1U53), (g) Pry1 (5ETE) and (h) pseudechetoxin (2DDA) are shown in the same orientation and the location of the two conserved Glu and His residues are marked red, indicating the CAP cavity. C CAP cavity formed by the conserved amino acid residues across CAP superfamily. The superposed structures of stecrisp (Trimeresurus stejnegeri; 1RC9), red; natrin (Naja atra; 1XX5), cyan; pseudechetoxin (Pseudechis australis; 2DDA) magenta and glioma pathogenesis-related 1 (GLIPR1) (Homo sapiens; 3Q2R), yellow showing key solvent exposed residues corresponding to His60, Glu75, Glu96 and His115 within the representative CAP domain. To superimpose CAP cavity, all the PDB structures were oriented and aligned based on structural similarity using PyMOL v2.1.0. CAP proteins across prokaryotes and eukaryotes are generally accepted as having a CAP domain containing a large ‘central cavity’, also known as the CAP cavity, active site or pocket site (Wilbers et al., 2018). Two variants of this cavity exist, one in which the CAPs have a CRISP-like cavity, which contains four key partially solvent-exposed amino acid residues present across the four CAP motifs: His from CAP1, Glu from CAP2, His from CAP3 and Glu from CAP4. These putative His-Glu pairs are present at the carboxyl terminal region of the CAP domain (Asojo and Koski, 2011). Based on crystal structures from reptile CRISPs, conserved residues corresponding to His60, Glu75, Glu96 and His115 form the CAP cavity (with sometimes the exception of Gln substituted at Glu75 position and Glu substituted at His60 position) (Fig. 3B and C) (Guo et al., 2005; Shikamoto et al., 2005). The large CAP cavity is stabilized by hydrogen bonding between His60 and His115 with the carboxyl oxygen atoms of Glu75 and Glu96 at the α3-helical strand and α6-helical strand, respectively (Shikamoto et al., 2005). The conserved histidine coupled with glutamic acid residues are oriented at the centre of the cavity and can directly complex with divalent cations, such as Zn2+, Mg2+ and Hg2+ (Henriksen et al., 2001; Wang et al., 2010; Darwiche et al., 2016; Darwiche et al., 2017). Moreover, the two histidine residues are solvent exposed and have been proposed to mimic protease active sites, as suggested by the ability of histidine to complex with the calcium-activated serine proteases-like activity in the cone snail CAP Tex31 (Milne et al., 2003). However, no further evidence exists to validate these findings. A similar His/Glu structural makeup is observed in the CAP domain from bacterial SCP domain-containing protein from the hookworm CAPs, Na-ASP-1 and Na-ASP-2 (Yamazaki et al., 2002; Guo et al., 2005; Shikamoto et al., 2005; Wang et al., 2005; Asojo, 2011; Darwiche et al., 2016). By contrast, the majority of the reported sequences of plant and parasite CAPs have the non-CRISP-like protein cavity and lack the histidine residue Asojo et al., 2018; Wilbers et al., 2018). The biological relevance of this variability remains undetermined. The hinge domain and ion channel regulatory domains (CRISP domain) The defining feature of CRISPs is the presence of the CRISP domain. It contains 10 conserved cysteine residues, which forms five intra-domain disulphide bonds. As indicated above, the CRISP domain can be further subdivided into two domains: the hinge domain, which is broadly seen in other members of the CAP superfamily, including GLIPR1, GAPR, CRISPLD1, CRISPLD2, PI15, PI16 and CLEC subfamilies, and a CRISP-specific ICR domain. The hinge domain comprises 20 amino acids and forms a ridged link connecting the CAP and C-terminal domains (in the case of the CRISPs, the ICR domain). The hinge is associated with the CAP domain at the carboxyl terminal residues—Pro160 and Tyr161. The rigidity of the hinge domain is imparted by the presence of two crossed disulphide bonds within its sequence, which prevents interactions between the CAP and ICR domains (Guo et al., 2005). Based on protein homology, including the positioning of cysteine residues, this structure is likely conserved across all hinge-containing CAPs. Members of the CRISP subfamily all contain a characteristic C-terminal domain of approximately 40 amino acids, most commonly referred to as the ICR domain. It contains a hydrophobic core with three α-helixes stabilized by a total of three disulphide bridges formed between Cys192–Cys210 and Cys201–Cys214, and eight amino acid loops connected by Cys183–Cys216 (Fig. 4) (Guo et al., 2005). The structural conformation of the C-terminal region of the ICR domain shares a high degree of similarity to the voltage sensitive potassium channel regulators ShK and BgK (Tudor et al., 1996; Dauplais et al., 1997; Gibbs et al., 2006). The ion channel regulatory activity of the CRISP subfamily will be discussed in detail below. Figure 4 Open in new tabDownload slide Schematic of the conserved cysteine residues in Stecrisp (PDB ID: 1RC9). Ribbon structure of CRISP showing the conserved eight disulphide bonds in mammalian CRISP proteins. Three disulphide bonds in the CAP domain are marked as red (Cys56–Cys134), magenta (Cys129–Cys145) and green (Cys73–Cys148) whereas five disulphide bonds in the CRISP domain are marked as yellow (Cys167–Cys174), blue (Cys170–Cys179), black (Cys192–Cys210), purple (Cys201–Cys214) and orange (Cys183–Cys216). Figure 4 Open in new tabDownload slide Schematic of the conserved cysteine residues in Stecrisp (PDB ID: 1RC9). Ribbon structure of CRISP showing the conserved eight disulphide bonds in mammalian CRISP proteins. Three disulphide bonds in the CAP domain are marked as red (Cys56–Cys134), magenta (Cys129–Cys145) and green (Cys73–Cys148) whereas five disulphide bonds in the CRISP domain are marked as yellow (Cys167–Cys174), blue (Cys170–Cys179), black (Cys192–Cys210), purple (Cys201–Cys214) and orange (Cys183–Cys216). Subfamilies within CAP Superfamily Proteins CRISPs The CRISPs are 27–29-kDa proteins that contain 16 conserved cysteine residues, which form eight intra-molecular disulphide bonds. CRISPs are found predominantly in vertebrates and their expression is enriched in the mammalian male reproductive tract and the venom of reptiles (Mochca-Morales et al., 1990; Yamazaki and Morita, 2004; Gibbs and O’Bryan, 2007). Most mammals possess three CRISP genes: CRISP1, CRISP2 and CRISP3, whereas some rodents contain an additional gene, Crisp4 (Fig. 5) (Jalkanen, 2005; Gibbs et al., 2008). CRISPs have been shown to participate in various aspects of mammalian fertilization, and due to the presence of the ICR domain have been widely studied in reptiles, but also in mammals, for their ion channel regulatory activities (discussed in detail below). Due to their expression bias in the male reproductive tract, CRISPs have gained special attention in the field of reproductive biology. Figure 5 Open in new tabDownload slide Phylogenetic analysis of vertebrate CRISP proteins showing the major lineages within the subfamilies. The phylogenetic tree was constructed using maximum likelihood (ML) analysis and shows that the CRISPs subfamily can be divided into four major lineages: CRISP1, CRISP2, CRISP3 and the venom CRISP (vCRISP). We note the presence of numerous polytomies is evident in the phylogenetic tree. The phylogenetic relationship shows the similarity, gene duplication events and nomenclature inconsistency across the subfamily and is discussed in detail in ‘Subfamilies within CAP superfamily proteins’. The complete annotated tree with a full description of methodology is available as Supplementary Fig. S1. Figure 5 Open in new tabDownload slide Phylogenetic analysis of vertebrate CRISP proteins showing the major lineages within the subfamilies. The phylogenetic tree was constructed using maximum likelihood (ML) analysis and shows that the CRISPs subfamily can be divided into four major lineages: CRISP1, CRISP2, CRISP3 and the venom CRISP (vCRISP). We note the presence of numerous polytomies is evident in the phylogenetic tree. The phylogenetic relationship shows the similarity, gene duplication events and nomenclature inconsistency across the subfamily and is discussed in detail in ‘Subfamilies within CAP superfamily proteins’. The complete annotated tree with a full description of methodology is available as Supplementary Fig. S1. In humans, CRISP1 is localized to the short arm of chromosome 6p21.3, along with the CRISP2 and CRISP3 genes, and is a reflection of the gene duplication event described above, as step 11 of the CAP evolutionary process. In mice, Crisp1–3 are located on chromosome 17B2, and an additional gene, Crisp4, is located on chromosome 1A3. As noted previously, identifying orthologues between species is more difficult than anticipated (Abraham and Chandler, 2017; Vicens and Treviño, 2018). For example, the gene sequence identity between mouse Crisp4 and human CRISP1 (59%) is greater than that between mouse Crisp1 and human CRISP1 (40%), suggesting that Crisp4 is the orthologue of human CRISP1 in the mouse (Jalkanen, 2005). Similarly, rat Crisp4 shares 69% and 91% sequence identity with human CRISP1 and mouse CRISP4, respectively (Nolan et al., 2006). As a result of this latter observation, several years ago the rat Crisp1 gene (NP_074050) was re-designated as rat Crisp4 (NP_001034482), and there is no rat Crisp1 gene per se. Many other examples of this confusion exist throughout the literature. Given the importance of CRISPs, notably in fertility, we have attempted to resolve the confusion described above through a phylogenetic analysis. The detailed methods of how the phylogenetic tree construction was carried out are described in Supplementary Fig. S1. Simplistically, all identifiable CRISP sequences were downloaded from National Centre for Biotechnology Information (NCBI: http://www.ncbi.nlm.nih.gov/) and UniProt databases (http://www.uniprot.org/). As there were inconsistencies in naming CRISPs, we strictly defined them by the presence of the four characteristic CAP motifs and the presence of 16 cysteine residues. Duplicate sequences were removed, and the remaining 222 sequences used to create a phylogenetic tree to reveal orthologues based on sequence. Our maximum likelihood-based tree reconstruction and analysis of CRISP gene sequence variation enables us to examine the phylogenetic relationships among CRISPs in various species (Fig. 5 and Supplementary Fig. S1). This analysis shows that within vertebrates there are four distinct lineages, designated here by the predominance of names within each clade: CRISP1, CRISP2, CRISP3 and reptile venom CRISPs (vCRISPs). Several polytomies are evident in the phylogenetic tree (Fig. 5 and Supplementary Fig. S1). Here we discuss each of these lineages in turn, highlighting the CRISP genes contained within each clade, identifying suspected instances of gene duplication, and incorrect nomenclature where appropriate. Our phylogenetic analysis shows that CRISP1 proteins, across multiple species, group together to form a well-supported lineage within the phylogenetic tree (62% bootstrap support). Interestingly, within this lineage there is a subclade (51% bootstrap support) that includes rodent CRISP4 proteins. These data indicate that rodent Crisp4 genes are actually the product of a gene duplication event of Crisp1. As indicated in the functional analysis, the duplicated sequences subsequently specialized to ultimately have a different function. Suffice to say, CRISP1 and CRISP4 sequences are derived from the same gene and the lines between what is a CRISP1 and what is a CRISP4 have been blurred during evolution. In addition, our phylogenetic analysis shows that the Crisp4 gene is not unique to mice (Vicens and Treviño, 2018). Our analysis identified CRISP4 protein in rodents from the Spalacinae family, such as Upper Galilee Mountains blind mole-rat (Spalax galili); the Cricetidae family, including the prairie vole (Microtus ochrogaster), Chinese hamster (Cricetulus griseus) and golden hamster (Mesocricetus auratus); the Ochotonidae family comprising the American pika (Ochotona); and the Dipodidae family including the lesser Egyptian jerbora (Jaculus jaculus) and the brown rat (Rattus norvegicus). It is important to note that several genes designated as CRISP1 fall outside the main CRISP1 clade, and their phylogenetic affinities are unresolved in our analysis (Supplementary Fig. S1). This includes CRISP1 from lizards (Anolis) and frogs (Xenopus), while CRISP1 in some monkeys (Callithrix jacchus, Ateles geoffroyi, Lagothrix lagotricha, Tarsius syrichta) exhibits a stronger homology to venom CRISPs (92% bootstrap support). CRISP2 forms the second lineage within the phylogenetic tree. The phylogenetic relationships within this clade align relatively well with the literature and databases, with a few exceptions. For example, mouse lemur CRISP1 is actually a CRISP2, or a gene duplication of CRISP2 as shown by this analysis. Interestingly, some CRISP2 sequences from goats (90% bootstrap support), and frogs (Xenopus) (92% bootstrap support), show a stronger homology to venom CRISPs than to the main CRISP2 lineage. CRISP3 forms the third lineage of the phylogenetic tree. CRISP3 does not form a single, well-supported lineage, as CRISP1 and CRISP2 do, but rather a series of poorly supported subclades, with no support for the relationships among the subclades. The main CRISP3 subclades include fish (86% bootstrap); birds/turtles (79% bootstrap); elephant/hyrax (100% bootstrap); Tasmanian devil (64% bootstrap) primates (77% bootstrap); cats/ferrets/panda (70% bootstrap); and rodents. Within the rodents, mouse CRISP1 and CRISP3 form a separate well-supported clade (100% bootstrap support), demonstrating a high sequence similarity between CRISP1 and CRISP3 in mice. This relationship indicates that mouse Crisp1 is actually a gene duplication of mouse Crisp3. Moreover, the analysis shows that rat CRISP3 is closely related to both mouse CRISP1 and CRISP3. Similarly, in prairie voles, CRISP1 appears to be a gene duplication of CRISP3. Interestingly, there is strong support (90% bootstrap support) for CRISP3 in ungulates (sheep, goats, horses) being more closely aligned with venom CRISPs than other CRISP3 sequences. As a result, researchers should not assume ungulate CRISP3 functional activity will be more comparable with other mammalian CRISP3 proteins as opposed to CRISP1 or CRISP2 functions. However, this homology to venom CRISPs is stronger (92% bootstrap support) in the lamprey. Reptilian venom CRISPs form the fourth distinct lineage within the CRISP phylogenetic tree (97% bootstrap support). This leads us to propose a new nomenclature for reptile CRISP—vCRISP (venom CRISP). Interestingly, CRISPs from the reptile Infraorder Platynota (Varanoidea; i.e. varanids and Gila monsters), the purported closest relatives of snakes (Streicher and Wiens, 2017), form a strongly supported sister lineage (92%) to vCRISPs. As mentioned previously, the vCRISPs also display a close homology to lamprey and Amazon molly CRISP3 (92% bootstrap support), Xenopus CRISP2 (92% bootstrap support), CRISP1 in some monkey species (C. jacchus, A. geoffroyi, L. lagotricha, T. syrichta) (92% bootstrap support), sheep, goat and horse CRISP3 (90% bootstrap support) and goat CRISP2 (90% bootstrap support). While the above analysis is intellectually satisfying and provides a framework for the direct comparison of functional data on orthologues versus paralogues, this analysis also indicates that the literature is muddled. In order to avoid compounding this confusion, below we will define the localization and function of CRISPs based on tissue expression using the published naming, but where appropriate have included the corrected nomenclature. Within mammals, male germ cells encounter CRISPs during all phases of development and maturation. CRISP2 is the only member of the CRISP subfamily expressed in mammalian testis, although as noted in sections below, other non-CRISP members of the CAP superfamily are expressed. CRISP2 was first identified as a major component of guinea pig sperm acrosome contents and thus named acrosomal autoantigen-1 (Hardy et al., 1988). Later, it was identified in mouse and rats as a testis-specific gene, Tpx-1, where its mRNA was shown to undergo a significant translational delay before the protein was incorporated into the sperm acrosome, connecting piece and the outer dense fibres of the sperm tail (Kasahara et al., 1989; O’Bryan et al., 2001). This localization is conserved in at least mouse, human and guinea pig (Hardy et al., 1988; O’Bryan et al., 1998, 2001; Jamsai et al., 2008b). Unlike CRISP1, the CRISP2 gene is not androgen-regulated, nor is the protein modified by glycosylation (Haendler et al., 1997). The significance of CRISP2 in mammalian fertility is discussed below. In mammals, sperm released from the testis are morphologically replete, but functionally immature and lack the capacity for progressive motility and fertilization. Sperm gain fertilization competence during their transition through the three distinct segments of the epididymis: caput (the proximal segment), corpus (the middle segment) and cauda (the distal segment) (Aitken et al., 2007). As the sperm are transcriptionally and translationally inactive, this transformation is dependent on post-translational protein modification, and protein shedding or acquisition from the epididymis (Brooks, 1982; Asano et al., 2010). It is during epididymal maturation that sperm encounter CRISP1 and/or CRISP4. In humans, a single CRISP is expressed in the epididymis, namely CRISP1. Within the rat epididymis, there is also a single CRISP, now correctly called CRISP4, but originally named CRISP1 (Roberts et al., 2008). Within mice, there are two epididymal CRISPs, CRISP1 and CRISP4. As discussed above, our phylogenetic analysis suggests that mouse Crisp4 arose as a gene duplication event in Crisp1. In accordance with the expression in mouse and rat, Crisp1 and Crisp4 expression in principal cells is regulated by androgens (Eberspaecher et al., 1995; Krätzschmar et al., 1996; Haendler et al., 1997; Jalkanen, 2005). Consistent with the possession of signal peptides, CRISP1 and CRISP4 are secreted into the epididymal lumen where they surround the transcriptionally silent sperm (Jalkanen, 2005; Nolan et al., 2006; Reddy et al., 2008; Gibbs et al., 2011). In mice and humans, CRISP1 is highly expressed in the caudal region of the epididymis but low levels are seen throughout the epididymis (Thimon et al., 2008; Dorus et al., 2010; Gibbs et al., 2011; Hu et al., 2018). In the mouse, expression of Crisp4 is highest in the initial segments (III–IV) of the caput epididymis, although low levels of expression are observed in cauda epididymis (Jalkanen, 2005). As shown in the rat, sperm are exposed to two variants of CRISP4 (D and E, called CRISP1 at the time of publication), which bind to the sperm surface but with different affinities (Jalkanen, 2005; Nolan et al., 2006; Gibbs et al., 2011). Structurally, the D and E isoforms differ minimally, such that a 203 dalton substitution is found at the N-terminal region of the E isoform (Roberts et al., 2007, 2008). The smaller D isoform (30 kDa) binds loosely to the sperm surface and is easily disassociated upon washing, whereas the larger E isoform is more tightly associated with the sperm plasma membrane and remains attached to the sperm head after sperm maturation in the female reproductive tract (Roberts et al., 2003; Busso et al., 2007; Roberts et al., 2008). The potential roles of CRISP1 and CRISP4 in mammalian fertility are discussed below. Unlike other mammalian CRISPs, CRISP3 has a wide expression pattern including in salivary gland, lacrimal gland, seminal vesicles, prostate, pancreas and the immune system (Reddy et al., 2008). The CRISP3 gene in mice and humans is androgen regulated and protein is produced in both unglycosylated and N-glycosylated forms (Haendler et al., 1997; Udby et al., 2002, 2005; Volpert et al., 2014). In humans, CRISP3 is a potential prostate cancer prognostic marker (Bjartell et al., 2007; Hoogland et al., 2011; Ribeiro et al., 2011). The clinical relevance of CRISP3 in pathologies and immune function is discussed below. CRISP3 is also found in high concentrations in ejaculates of several species, including horses, (~21 ng/ml) where its concentration, or bioactivity, in semen has been correlated with higher first cycle pregnancy rates following artificial insemination (Novak et al., 2010; Waheed et al., 2015; Usuga et al., 2018; Restrepo et al., 2019). The role of CRISP3 in seminal plasma and fertility will be discussed in detail below. Collectively, this strong association between CRISPs and various aspects of fertility and their positive selection has led to the hypothesis that although CRISPs may not be essential for male fertility, they confer a benefit upon sperm function in situations of sperm competition or in the regulation of the immune system in the female reproductive tract wherein sperm are immunologically foreign. This hypothesis has not been directly tested. GLIPR proteins Members of the GLIPR subfamily of proteins possess a signal peptide followed by a CAP domain, a hinge region and a subgroup defining C-terminal extension (Ren et al., 2006; Gibbs et al., 2010). The C-terminal extension contains 13 cysteine residues, 12 of which are conserved with the CRISP subfamily (Gibbs et al., 2008, 2010). GLIPR1 was originally identified and characterized in human glioblastoma as a ‘human related to testes-specific, vespid, and pathogenesis protein 1’ (hRTVP1) (Murphy et al., 1995; Rich et al., 1996). In the mouse, it was identified as a p53 target gene with proapoptotic activity (Ren et al., 2002). Genome-wide sequence analyses have identified three human GLIPR genes (GLIPR1, GLIPR1L1 and GLIPR1L2) and four mouse genes (Glipr1, Glipr1l1, Glipr1l2 and Glipr1l3). All are clustered within the p53-regulated gene cluster on chromosome 12q21 and chromosome 10D1, respectively (Ren et al., 2006). Mouse GLIPR1L1 shares 44% sequence identity with human GLIPR1L1 and macaque GLIPR1L1 (also called MAK248) and 48% sequence identity with bovine GLIPR1L1. Of note, in contrast to mouse and bovine GLIPR1L1 which is expressed in the testis and incorporated into the apical pole and connecting piece of the elongated spermatids, macaque GLIPR1L1 is exclusively expressed in the epididymis and linked by a glycosylphosphatidylinositol (GPI) anchor to the sperm plasma membrane overlying the acrosome (Yudin et al., 2002; Gibbs et al., 2010; Caballero et al., 2012). In all species examined to date, GLIPR1L1 appears to play a facilitative role in sperm-oocyte binding (Gibbs et al., 2010; Caballero et al., 2012; Gaikwad et al., 2019). The role of GLIPR1L1 in mammalian fertility will be described in detail in sections below. Interestingly, GLIPR1L2 has a glutamate-rich domain (ERD) C-terminal domain that contains 32 glutamate residues (Ren et al., 2006), the function of which is unknown. Other proteins containing a C-terminal ERD region, however, have been shown to mediate structural stability during protein–protein interactions, suggesting that the ERD region is a protein-interacting domain (Lee et al., 2008). GLIPR1 and GLIPR1L2 are expressed in diseased human tissues including prostate, bladder, kidney and bone marrow (Ren et al., 2006). Analysis of expressed sequence tag expression profile suggests that GLIPR1L3 (ENSMUSG00000112611) is found only in mouse and it arose via duplication of the Glipr1l1 gene (Ren et al., 2006; NCBI Resource Coordinators, 2016) (82% sequence identity) (Gibbs et al., 2008). Currently, there is no functional data on GLIPR1L3 although mRNA expression data indicate it is expressed in the testis (NCBI Resource Coordinators, 2016). GAPR proteins The GAPR protein subfamily is also referred to as GAPR-1 and ‘Homo sapiens chromosome 9 open reading frame 19’ (C9orf19). This CAP subfamily has the highest sequence homology with plant Pr-1 proteins and is considered as the most evolutionarily primitive member of the superfamily. They contain a single-CAP domain, which evolved during Steps 2–4 (Abraham and Chandler, 2017). There is one GAPR gene in humans and one in mice. GAPR proteins are often characterized by their small molecular mass (14–17 kDa) and resistance to proteases (Olrichs et al., 2014). They lack a signal peptide suggesting they serve intracellular functions (Eberle et al., 2002). The GAPR gene was originally characterized in human as C9orf19 gene and linked to the aetiology of hereditary inclusion body myopathy (IBM2) (Eisenberg et al., 2002). IBM2 belongs to a group of rare autosomal recessive disorders and is characterized by muscle weakness, especially of the quadriceps (Huizing and Krasnewich, 2009). In human, the GAPR gene is localized on chromosome 9p12-p13 and is highly expressed in peripheral blood leukocytes and lungs (Eberle et al., 2002; Eisenberg et al., 2002). In mice, Gapr is localized on chromosome 4B1 and is predominantly expressed in the kidney, pancreas, lung and uterus (Eberle et al., 2002). The biological function of GAPR proteins is largely unknown; however, a key role in the immune system of mammals is now emerging. The role of GAPR proteins in autophagy was reported by Shoji-Kawata and colleagues (Shoji-Kawata et al., 2013). Autophagy is a highly conserved cellular ‘recycling’ mechanism, which maintains cellular integrity by degrading old intracellular proteins and damaged organelles (Yorimitsu and Klionsky, 2005; Lucin and Wyss-Coray, 2013). Specifically, GAPR was identified as a putative binding partner of an 18 amino acid peptide from beclin1, a protein which enhances autophagy by targeting the lysosomal degradation pathway (Shoji-Kawata et al., 2013). GAPR was shown to sequester beclin1 at the Golgi apparatus and limit its function, suggesting that GAPR negatively regulates autophagy (Shoji-Kawata et al., 2013). Furthermore, GAPR has been associated with amyloid-related diseases and has the ability to form amyloid structures (Sheng et al., 2019). GAPR was localized in the microdomains of the Golgi complex, which are highly enriched in cholesterol and sphingolipids, which are important in amyloid regulation in vitro (Fantini and Yahi, 2010; van der Meer-Janssen et al., 2010). GAPR has an affinity for negatively charged lipids, and studies have shown a time-dependent correlation between formation of amyloid-like fibrils, potentially suggesting a role of the CAP domain in amyloid-like oligomerization that may be of relevance across the superfamily (Olrichs et al., 2014; Sheng et al., 2019). CRISPLD1 proteins CRISPLD1 is also known as ‘CAP and LCCL domain containing protein-1’ (CAPLD1), Cocacrisp, CRISP10 and LCRISP1 (Gibbs et al., 2008)—although we note it is not a CRISP. Structurally, CRISPLD1 proteins contain CAP and hinge domains, and a subfamily-defining C-terminal LCCL domain (Gibbs et al., 2008). The LCCL domain is named after its sequence homology to Limulus factor C, cochlear protein Coch-5b2 and late gestation lung protein Lgl1 (Liepinsh et al., 2001). The LCCL domain is 100 amino acids in length and contains a conserved histidine residue in the signature motif [YxxxSxxCxAAVHxGVI] in its C-terminal region (Trexler et al., 2000). There is one CRISPLD1 protein in humans and one in mice. Circular dichroism spectrum analysis of the LCCL domain from recombinant human CRISPLD1 revealed a high β-sheet content (54%) (Trexler et al., 2000). Most, but not all, LCCL domains have four highly conserved cysteine residues, which form two intra-domain disulphide bonds (Cys8-Cys28 and Cys24–48) linking the α-helix and β-sheet at the N-terminal of the LCCL domain (Trexler et al., 2000). The LCCL domain has been proposed to have structural and anti-microbial functions, along with roles in protein, lipid and carbohydrate binding (Liepinsh et al., 2001; Dessens et al., 2004; Pradel et al., 2004; Tremp et al., 2017). In humans, CRISPLD1 is localized on chromosome 8q21.11 and in mice, on chromosome 1A4. In humans, mutations in the LCCL domain (P25S, V40G, G62E and I83N) result in misfolded protein, potential loss-of-function and deafness autosomal dominant non-syndromic sensorineural disorder 9 (DFNA9) (de Kok et al., 1999; Trexler et al., 2000; Liepinsh et al., 2001). The LCCL domain has also been implicated in eye disease, where it is upregulated in patients with glaucomatous trabecular meshwork, compared to controls (Trexler et al., 2000; Picciani et al., 2007). A single nucleotide polymorphism (SNP) in CRISPLD1, rs12115090, A>C allele in carrier patients with cardiovascular diseases, has been shown to result in a significantly reduced risk of clopidogrel drug resistance, suggesting an association of CRISPLD1 and stem cell differentiation (Wang et al., 2018). However, the exact mechanism behind this phenomenon remains unknown. A recent study using a mouse model identified CRISPLD1 in the extracellular matrix of articular cartilage involved in osteoarthritis pathophysiology (Wilson et al., 2016). CRISPLD2 proteins The CRISPLD2 proteins are also referred to as CAPLD2s. Like CRISPLD1, CRISPLD2 contains a signal peptide, a CAP domain, a hinge domain and a unique C-terminal extension consisting of 260 amino acid residues of tandem LCCL domains (Gibbs et al., 2008). There is one CRISPLD2 protein in humans and one in mice. In humans, CRISPLD2 is localized on chromosome 16q24.1 and in mouse chromosome 8E1. Studies have suggested the involvement of CRISPLD2 in kidney and lung development and mutations in CRISPLD2 have been controversially linked to the aetiology of craniofacial morphogenesis (Quinlan et al., 2007; Zhang et al., 2015; Zhang et al., 2016). For example, data from a Chinese population supported the association between CRISPLD2 gene mutations and syndromic cleft lips, with or without cleft palate (nsCL/P), whereas this associated was not seen in a European population (Chiquet et al., 2007; Shi et al., 2010; Girardi et al., 2011; Mijiti et al., 2015). In addition, CRISPLD2 has been associated with chronic bronchopulmonary dysplasia (BPD). Crispld2 expression was significantly downregulated in an animal model of BPD, homozygous deletion of Crispld2−/− in mice leads to embryonic lethality, and heterozygous Crispld2+/− mice have features representative of BPD (Nadeau et al., 2006; Lan et al., 2009). The knockdown of Crispld2 in human airway fibroblasts led to a significant increase in expression of the proinflammatory mediators interleukin (IL) 8, chemokine (C–C motif) ligand 2 (CCL2) and IL6, suggesting that CRISPLD2 has anti-inflammatory properties that can reduce pro-inflammatory signalling by lung epithelial cells through mesenchymal–epithelial interactions (Zhang et al., 2016). Further, CRISPLD2 expression was significantly downregulated (−2689 relative fold change) in patients with Gaucher disease, a rare inherited metabolic disease (Ługowska et al., 2019). This finding has yet to be investigated in detail. PI15 proteins PI15 was first identified as a 25-kDa novel trypsin inhibitor, also referred to as p25TI, sugarcrisp and CRISP8 (Koshikawa et al., 1996; Lambeth et al., 2014). Again, it is not actually a member of the CRISP subfamily. PI15 is found in a range of mammals including human, chimpanzee, monkey, cow, horse, dog, mouse and rat. Within humans and mice, there is one PI15 gene. Structurally, PI15 proteins contain a signal peptide, a CAP domain, a hinge domain and a 12 amino acid C-terminal extension. In human, PI15 is located on chromosome 8q21.11 and is predicted to produce a mature 198 amino acid protein following cleavage at a furin-like protease cleavage site (Yamakawa et al., 1998). Within mice, Pi15 is localized on chromosome 1A4. Immunochemistry data from human and mouse tissues identified PI15 production in the prostate, mammary and salivary glands, thyroid, skeletal muscle, smooth muscle, heart and ovary (Smith et al., 2001). Mouse PI15 has been identified as a potential regulator of podocyte development in the kidney. PI15 expression is highly upregulated in kidney at embryonic day (E)18.5, mimicking forkhead box protein C2 (FOXC2)—a key transcription factor required for podocyte development, making it a potential biomarker for kidney function and disease (Takemoto et al., 2006). PI15 has also been implicated in craniofacial development such that upregulation of PI15 in chicken embryo leads to cleft beak, analogous to human cleft lip (Nimmagadda et al., 2015). Previously, PI15 expression has been found in the lining of human atherosclerotic aortic wall and in mouse hyperlipidemic aorta (Hägg et al., 2009). Furthermore, the downregulation of PI15 in rat aorta resulted in increased protease activity leading to rupture of the internal elastic lamina, suggesting anti-inflammatory properties of PI15 (Falak et al., 2014). It has also been suggested that PI15 plays a role in the development of infectious Chlamydia trachomatis, in which the pathogen recruits low concentrations of host PI15 to regulate chlamydial protease-like activity factor (CPAF). Nonetheless, the exact mechanism by which PI15 activates CPAF largely remains unexplored (Prusty, Chowdhury, Gulve, & Rudel, 2018). It is suggested that PI15-CPAF forms a stable multimeric complex within the inclusion lumen via the Golgi-associated vesicles. By contrast, over-expression of PI15 has an inhibitory effect on the CPAF protease activity due to delayed chlamydial growth, suggesting that PI15 could potentially be a protease inhibitor (Prusty et al., 2018). PI16 proteins PI16 is also known as cysteine-rich protease inhibitor (CRIPI), PSPBP and microseminoprotein-β (MSMB) and was referred to as CRISP9 in some public databases, although it is clearly not a member of the CRISP subfamily (Reeves et al., 2005; Frost and Engelhardt, 2007; Hazell et al., 2016). In human, PI16 is localized to chromosome 6p21.2 and encodes one PI16 protein. In mice, it is localized on chromosome 17B1 and encodes one PI16 protein. PI16 protein contains a signal peptide, a CAP domain, a hinge domain and a ~264 amino acid C-terminal extension, which is predicted to be unstructured (Regn et al., 2016). PI16 was identified during a screen of a mouse cardiac cDNA library for secretory proteins and as a paracrine factor with potential cardiac function, where PI16 levels were highly upregulated (Frost and Engelhardt, 2007). The exact role of PI16 is unknown; however, a genome-wide association study suggested a correlation between SNPs in the CAP domain of PI16 (rs1405069) and elevated chemerin plasma levels and that PI16 and chemerin levels are closely associated with heart disease (Bozaoglu et al., 2010; Rodríguez-Penas et al., 2015; Zhang et al., 2015; Regn et al., 2016). PI16 is proposed to regulate the myocardium by inhibiting proteolytic activation of pro-chemerin, which subsequently prevents chemerin-dependent activation of the G protein-coupled receptor CMKLR1, which in turn inhibits pro-inflammatory pathways (Regn et al., 2016). PI16 also has a role in inflammation and immune regulation. In response to host infection, PI16 expression was upregulated and localized to the surface of CD4+/CD25+ regulatory T cells (Treg cells) (Nicholson et al., 2012; Hope et al., 2019). The PI16+ Treg cells are a subset of FOXP3+ memory Treg cells, which are capable of migrating towards the chemokines CCL17 and CCL20 at sites of inflammation (Nicholson et al., 2012), and this suggests they regulate pro-inflammatory responses (Miyara et al., 2009; Sadlon et al., 2010; Nicholson et al., 2012). This finding is of interest also because CCL17 is recognized by the CCR4 chemokine receptor on skin-homing T cells, and studies have hypothesized the association between PI16 and cutaneous homing of T cells (Homey et al., 2002) where PI16 was found in association with CD8+ T cells in patients with acute graft versus host disease and controls (Lupsa et al., 2018). It has also been proposed that PI16 is a potent inhibitor of matrix metalloproteinase-2 (MMP-2) activity of relevance to skin homeostasis (Hazell et al., 2016). MMP-2 is an extracellular protease, which plays a significant role in cell growth, migration, invasion, inflammation, angiogenesis, cancer development and ischemia–reperfusion injury (Cheung et al., 2000; Puyraimond et al., 2001; Schmalfeldt et al., 2001; Wang et al., 2002; Klei et al., 2004; Bauvois, 2012; Hazell et al., 2016). CLEC proteins The CLEC proteins are also referred to as mannose receptor-like proteins, and CAP domain- and CTL domain-containing proteins. They contain a signal peptide and a CAP domain, followed by a hinge domain and a subfamily defining 115–130 amino acid C-terminal extension known as the C-type lectin-like domain (CTLD) (Drickamer, 1999; Drickamer and Dodd, 1999; Dodd and Drickamer, 2001). The CTLD domain is also known as the carbohydrate recognition domain. Originally, the CTLD domain was identified as a Ca2+-dependent carbohydrate recognition and binding domain in animal lectins (Drickamer, 1988, 1999). The CTLD domain contains four cysteines, which form a characteristic ‘loop-in-loop’ fold stabilized by a two highly conserved disulphide-bond bridges (Zelensky and Gready, 2005). In human, there are three CLEC subfamily members. CLEC18A and CLEC18B are located on chromosome 16q22.1, and CLEC18C is located on chromosome 16q22.3. A single mouse gene is located on chromosome 8E1. In humans, the CTLD domain of CLEC18A is speculated to have high specificity and affinity for glycoproteins and to play a role in glycolipid transport mechanism (Huang et al., 2015). In addition, the CLTD domain has been implicated in immune response regulation, inflammation and autoimmunity (Ezekowitz et al., 1990; Lee et al., 2002; Schneiter and Di Pietro, 2013; Olrichs et al., 2014; Huang et al., 2015). In humans, the CLEC18 proteins (CLEC18A, CLEC18B and CLEC18C) are highly expressed in peripheral blood cells and are upregulated during monocyte differentiation processes (Huang et al., 2015). CLEC proteins have been speculated to bind glycolipids and promote glycolipid transport (Huang et al., 2015). Moreover, they have been suggested to inhibit pathogen invasion by extracting sterol components from the pathogens, including members of flaviviruses, in the endoplasmic reticulum and Golgi apparatus (Huang et al., 2015). As shown by the work of Choudhary et al., the ability to bind sterols is almost certainly mediated by the CAP domain (Choudhary and Schneiter, 2012). In Caenorhabditis elegans, members of the CLEC proteins, CLEC39 and CLEC49, are homologous to human CLEC18A/CLEC18B and have been suggested to play a significant role in immune responses, such that Clec39 and Clec49 are highly upregulated in response to fungal and bacterial invasion (Harcus et al., 2009; Miltsch et al., 2014). Further, the survival rate and fertility of C. elegans was significantly decreased in Clec39−/− and Clec49−/− worms when infected with Serratia marcescens, suggesting a role in immunity and bacterial infection (Miltsch et al., 2014). In humans, CLEC18A levels were found to be significantly increased in patients with chronic hepatitis C virus (HCV) and positively associated with viral load, cryoglobulin and C4 levels in patients with mixed cryoglobulinemia syndrome, suggesting its potential utility as a biomarker of HCV-associated mixed cryoglobulinemia syndrome (Liao et al., 2018). In addition to liver disease, patients with HCV infection are also reported to have various HCV-related extrahepatic manifestations (HCV-EHMs) (Sansonno et al., 2007), with mixed cryoglobulinemia syndrome being the most common HCV-EHM disease (Scotto et al., 2006). The venom Ag proteins The venom allergen proteins (Ag), previously known as antigen 5 (Ag5) proteins, are a founding subfamily in the CAP superfamily (Gibbs et al., 2008). The venom Ag were one of the earliest members of the CAP superfamily to evolve and are examples of the Step 2–4 proteins described in relevant section above. They are found in fire ants and numerous wasp species, but not in mammals. They contain a single CAP domain free of other domains. Interestingly, in certain members of this subfamily (parasitic wasp—Ag3, Ag5; Jewel wasp—Ag5-like, Ag3-like; Red fire ants—Ag3 and Leafcutte bee—Ag3-like), the CAP defining cysteines are present in the N-terminal signal peptide, and as such they are not found in the mature protein (Abraham and Chandler, 2017). As a consequence, they do not contain the characteristic CAP cavity described above, although they have significant identity at the gene sequence level. The Ag proteins are found in high levels in wasp venom and include Dol m 5 in white-faced hornet (Dolichovespula maculata), Vesp c 5 in the European hornet (Vespa crabro), Pol a 5 in the paper wasp (Polistes annularis), Ves v 5 in the yellow jacket (Vespula vulgaris) and Sol I 3 protein in fire ants (Solenopsis invicta) (Hoffman, 1993; Lu et al., 1993; Tomalski et al., 1993; Monsalve et al., 1999; King and Spangfort, 2000). These proteins are strongly antigenic in humans and induce an allergenic response (King and Spangfort, 2000; Henriksen et al., 2001; Cifuentes et al., 2014; Justo Jacomini et al., 2014). In tsetse flies, Ag proteins have been hypothesized to suppress the immune response in the gut of the host (Li et al., 2001). However, this hypothesis remains untested. Nonetheless, Ag5 is one of the major allergens characterized in vespid venom insects such as hornets, yellow jacket and paper wasps (King et al., 1978; King and Spangfort, 2000). The Ag5 proteins induce a strong allergic response in affected individuals (King et al., 1978; King et al., 1983; Hoffman, 1993), which involved both IgG and IgE components (Monsalve et al., 1999; Bohle et al., 2005). In Drosophila melanogaster, 26 Ag proteins have been identified, of which 70% showed enriched expression in the testis, suggestive of roles in fertility (Kovalick and Griffin, 2005). It is tempting to hypothesize that Ag proteins in insects could have a similar function in fertility and immunity to that of mammalian CAP superfamily proteins. The bacterial SCP domain-containing proteins The bacterial SCP domain-containing proteins are made up of single CAP domains and are examples of step 1 CAP. It is worth noting that these proteins are not just found in bacteria, but also extend to parasitic helmiths, and they go by a variety of alternative names including anclyostoma-secreted proteins, activation-associated secreted proteins (ASPs), ASP-like protein, venom allergen-like (VAL) proteins, venom allergen-homologous (VAH) proteins and venom allergen-proteins (VAP) (Frank et al., 1996; Gao et al., 2001; Gibbs et al., 2008; Saverwyns et al., 2008; Visser et al., 2008; Cantacessi et al., 2009; Stroehlein et al., 2016; Wilbers et al., 2018). The SCP domain-containing proteins have been extensively characterized and divided into three groups: clade III (nematode, Spirurida and parasitic nematode round worms, Ascaridida); clade IV (plant-parasitic nematode, Tylenchida); and clade V (hookworms, Strongylida and microbivorous nematode, Rhabditida) (Cantacessi et al., 2009). Parasitic worms (helminths) have been extensively studied for their ability to survive in hosts through manipulation of the host immune response (Goverse and Smant, 2014; Maizels and McSorley, 2016). CAP proteins have been implicated in this process. Specifically, VAL expression is significantly upregulated during the parasitic phases of helminth life cycles, both in plants and animals (Hewitson et al., 2009; Lozano-Torres et al., 2012; Heizer et al., 2013; Sotillo et al., 2014; Hunt et al., 2016), and they are associated with the parasite surface (Hewitson et al., 2011). Further, proteomic analysis of Heligmosomoides polygyrus, an intestinal helminth parasite in rodents and humans, revealed a high level of immunomodulatory components, including VAL-proteins and ASP-proteins, during infection (Hewitson et al., 2011). A similar situation occurs in humans infected with Strongyloides species, including Strongyloides stercoralis, where a number of bacterial SCP domain-containing proteins are significantly upregulated during the parasitic stage as compared to free-living lifestyle [for a complete list refer to reference (Hunt et al., 2016)]. Within clade III, the dog hookworm (a nematode) Ancylostoma caninum expresses a variety of the SCP domain-containing proteins including Ac-ASP-1 (double CAP-domain ASP) and Ac-ASP-2 (C-type single CAP-domain ASP) (Gibbs et al., 2008; Cantacessi et al., 2009; Cantacessi and Gasser, 2012). In adult A. caninum, ASPs are also called neutrophil inhibitor factor and they inhibit both adhesion of activated human neutrophils to endothelial cells and the release of hydrogen peroxide from activated neutrophils (Moyle et al., 1994). In addition, similar to other members of the CAP superfamily, including CRISPs, GAPR1, Pr1 and CLEC, the bacterial SCP domain-containing proteins have sterol binding activity (Choudhary and Schneiter, 2012; Choudhary et al., 2014; Asojo et al., 2018). SCP domain-containing proteins produced by plant-parasitic nematodes (Clade IV) provide additional evidence of a role for CAP domains in host immune suppression. For example, during the parasitic phase within its plant host, the nematode Heterodera glycines expresses two venom allergen-like proteins, hg-vap-1 and hg-vap-2, in the subventral oesophageal gland cells (Gao et al., 2001). Further, Meloidogyne incognita, a root-knot nematode, expresses two ASP proteins, meloidogyne secretory protein-1 (Mi-MSP-1) and Mi-VAP-2, which are highly expressed in pre-parasitic and parasitic second-stage juveniles (J2) (Ding et al., 2000; Cantacessi and Gasser, 2012). Both proteins play a role as activators and suppressors of host immune response in plants (Ding et al., 2000; Wang et al., 2007). Collectively, these data indicate the significant role of the bacterial SCP domain-containing proteins in suppression of the immune system. The structure of recombinant HpVAL-4, a clade V SCP domain-containing protein from the parasitic hookworm Heligmosomoides polygyrus bakeri, has been reported (Hawdon et al., 1996; Borloo et al., 2013; Asojo et al., 2018). HpVAL-4 contains a caveolin-binding motif (CBM) and a palmitate-binding cavity suggesting a comparable cholesterol and fatty acid-binding ability to Pry1 and tablysin-15 in vitro (Ma et al., 2011; Asojo et al., 2018). Data also suggest that multiple bacterial SCP domain-containing proteins show preferential expression in the parasitic form or free-living form of the same species: an interesting example of this being the Strongyloides nematodes, which belong to clade V of the SCP domain-containing proteins and which can adapt to parasitic or free-living life cycle (Hunt et al., 2016). The ASPs are another group of bacterial SCP domain-containing protein produced by hookworms. ASPs are unusual in that several members contain multiple CAP domains in tandem: the C-type single domain ASPs, double-domain ASPs (two SCP-domain) and N-type single-domain ASPs (Geldhof et al., 2003). ASPs are abundantly produced by infective third-stage larvae (L3) in dogs, where they have been shown to play an important role in the transition from the free-living to parasitic life-cycle during invasion of the host (Hawdon et al., 1999; Datu et al., 2008). Finally, in the trematodes (Schistosoma mansoni), 13 SCP proteins (SmVAL1–SmVAL13) have been identified. Twelve contain a single CAP domain, and one member contains two CAP domains (Chalmers et al., 2008; Cantacessi and Gasser, 2012). To date, their roles are unexplored. However, after infecting their snail host, S. mansoni begin to transform from the miracidium to parasitic sporocyst stage (Basch and DiConza, 1974; Lodes and Yoshino, 1989), which includes the production of high quantities of SmVAL proteins. SmVAL proteins have been shown to play a role in stress responses, proteolysis/inhibition, as an antioxidant and in detoxication, and in immune modulation, suggesting they may play a role in protecting the parasite against host defense mechanisms (Wu et al., 2009). Of note, when the three Onchocerca volvulus (a nematode which causes blindness) C-type single-domain ASPs (Ov-ASP-1, Ov-ASP-2 and Ov-ASP-3) where injected into mice they induced angiogenesis (Lizotte-Waniewski et al., 2000; Tawe et al., 2000). Ov-ASPs also show a characteristic expression pattern during parasite development, such that Ov-ASP-1 is abundant in second-stage larvae (L2s), Ov-Asp-3 is specific to the third-stage larvae (L3s) and Ov-Asp-2 is present in all developmental stages (Tawe et al., 2000; Cantacessi and Gasser, 2012). These data suggest that SCP domain-containing proteins may contribute to multiple aspects of parasite host invasion. The Pr-1 proteins The Pr-1 proteins are proposed to have evolved from the bacterial SCP proteins and are characterized by the presence of a CAP domain, containing the four CAP motifs, and two to four Pr-1 specific cysteine residues within the CAP domain (Abraham and Chandler, 2017). The Pr-1 proteins are plant-specific and are highly upregulated in response to pathogenic infection caused by bacteria, viruses and fungi (Breen et al., 2017). This phenomenon was first demonstrated by Loon Van and Kammen Van who infected the leaves of tobacco plant (Nicotiana tabacum) with tobacco mosaic virus and showed that multiple Pr-1 proteins accumulated, up to 10% of the total protein in the infected leaf (Van Loon and Van Kammen, 1970; van Loon, 1985; Cornelissen et al., 1986). This suggested a defensive role. There is also evidence suggesting roles for Pr-1 proteins in plant development, where expression is increased in leaves of adult flowering plants and concentrated in the sepals of the developing flowers (Fraser, 1981; Lotan et al., 2007). Plant proteomic analysis identified a variety of Pr-1 proteins in the leaves and seedlings of Arabidopsis thaliana (Arabidopsis), the hypocotyls of Helianthus annuus (sunflower) and roots of Pisum sativum (peas) (Chaki et al., 2009; Begara-Morales et al., 2013; Takahashi et al., 2015). Pr-1 proteins, including Pr-1A, Pr-P and thaumatin-like protein E2, were found to undergo protein tyrosine nitration under induced abiotic stress (Takahashi et al., 2016). PR-1B and thaumatin-like protein E2 have been shown to have antimicrobial functions, including enzymatic activities to prevent microbial growth (Leah et al., 1991; Elvira et al., 2008). Pr-1 proteins have also been suggested to play a role in the surveillance mechanism in the plant immune system (Edreva, 2005). Together, these data suggest multiple functions of plant Pr-1 in the development and immune regulation. The fungal PRY proteins While initially classified as a member of the Pr subfamily, a more in-depth phylogenetic analysis revealed that fungal PRY proteins form a distinct clade within the CAP superfamily, which evolved independently (Gibbs et al., 2008; Choudhary and Schneiter, 2012; Abraham and Chandler, 2017). Of likely relevance across the CAP superfamily, the Saccharomyces cerevisiae proteins PRY1 and PRY2 were the first CAPs to be characterized as sterol binding proteins with the ability to export sterol in vivo and bind to cholesterol in vitro (Choudhary and Schneiter, 2012). A recent study also suggests that PRY1 bind to sterols and fatty acids at different binding sites in the CAP domain (Darwiche et al., 2017). Computational modelling of PRY1 identified the region involved in sterol binding and showed it contains a flexible loop, similar to CBM, which also regulates the export of cholesteryl acetate in vivo (Choudhary et al., 2014). Consistent with this observation, truncating the PRYI CAP domain led to impaired lipid export in yeast (Choudhary and Schneiter, 2012). Structural analysis of the PRY1 CAP domain also revealed that sterol binding is facilitated by magnesium binding regions within the CAP domain (Darwiche et al., 2016). Based on this, and the fact that the sterol binding ability is conserved in a mammalian CAP (CRISP2) (Choudhary and Schneiter, 2012), it is tempting to hypothesize that CAP domains across the superfamily can accommodate binding to diverse lipids (as discussed with other members of the protein family including GAPR-1, Pr-1, CLEC and the bacterial SCP domain-containing proteins). The Role of CAP Superfamily Protein in Male Fertility and Pathologies The CAP superfamily proteins are strongly associated with numerous aspects of mammalian fertility. Members of the CAP proteins are expressed in the testes during spermatogenesis, epididymal CAP proteins are involved in sperm maturation and potentially involved in sperm–egg fusion and accessory gland-derived CAPs are potentially involved in modulating the female reproductive tract to optimize male fertility. In this section, we will discuss the roles and associations of CAP superfamily proteins during the various phases of sperm development and function in the hope of providing a holistic view of their importance in mammalian fertility. The role of CAPs in spermatogenesis The testes are the sites of spermatogenesis, a process during which germ cells differentiate into sperm. In mammals, Crisp2 and Glipr1l1 are the only members of the CAP superfamily characterized as having roles in the testis, although an examination of expression databases suggests that others, including GLIPR1L3 (ENSMUST00000220085), PI16 (ENSMUST00000114701), CRISPLD1 (ENSMUST00000095075) and CRISPLD2 (ENSMUST00000034282), are also expressed in the testes in various species (Maeda et al., 1999; Gibbs et al., 2010). The assignment of the name CRISP2 has been relatively consistent across species at least in regard to localization and functional data, if not strictly to sequence annotations (Fig. 5). As described above, CRISP2 is incorporated into the sperm acrosome, along the sperm tail as part of the outer dense fibres, and the connecting piece at the junction between the head and tail. Data from Crisp2 knockout mice indicate that the incorporation of CRISP2 into the acrosome is functionally relevant. Specifically, Crisp2 knockout mice have a severely stunted ability to manifest the acrosome reaction in response to progesterone (Lim et al., 2019). Moreover, sperm from Crisp2 null mice have a stiff mid-piece, which severely compromises motility and fewer numbers of sperm are able to penetrate oocytes (Lim et al., 2019). As a result, Crisp2 null mice are subfertile (Lim et al., 2019). A second knockout model indicated that Crisp2 null sperm had a compromised ability to fertilize in vitro under demanding conditions (Brukman et al., 2016). Whether the latter is due to a compromised acrosome function and/or motility defects or a fertilization-specific defect is impossible to dissect out (Brukman et al., 2016; Lim et al., 2019). Early data also suggested that CRISP2 may be involved in anchoring germ cells to Sertoli cells within the seminiferous epithelium via the N-terminal region of the CAP domain (Maeda et al., 1998). Data from the Crisp2 null mouse lines indicate that this role is not, however, critical and that the loss of CRISP2 did not elevate germ cell sloughing or affect daily sperm output (Lim et al., 2019). In retrospect, the assignment of Sertoli-germ cell adhesive properties may have been incorrectly applied based on the differences in the secretory pathways in haploid male germ cells and the Jurkat cells used in the study (Maeda et al., 1998; Lim et al., 2019). These data do, however, indicate that the N-terminal 101 amino acids of the CRISP2 CAP domain have cell-adhesive properties (Maeda et al., 1999) and may thus be relevant in other contexts, or to other CAP proteins. Like many of the reptile CRISPs (see below), CRISP2 can regulate ion channels present in sperm, consistent with a role in regulating sperm motility (Harper et al., 2004; Koppers et al., 2011). The CRISP2 CRISP domain can regulate Ca2+ flow through ryanodine receptor channels (RyR1 and RyR2) (Gibbs et al., 2006). Data also indicate that CRISP2 can bind to, and potentially regulate, the cation channel of sperm 1 (CATSPER1) subunit of the CatSper ion channel, which is necessary for normal sperm motility. This has not, however, been directly tested although dysregulated intracellular Ca2+ levels during capacitation have been observed in Crisp2−/− sperm (Ren and Xia, 2010; Brukman et al., 2016; Lim et al., 2019). Mounting data also suggest a significant role for CRISP2 in human male fertility that is, by and large, consistent with the mouse data. Data indicate that CRISP2 mRNA stability is negatively regulated by micro RNA (miRNA)-27a and mi-RNA-27b binding to its 3′-untranslated region (Zhou et al., 2015, 2016) and that there is a significant association between CRISP2 protein content and the concentration of the two miRNAs in sperm from patients with asthenoteratozoospermia (abnormal sperm motility and morphology). In concordance, higher Crisp2 mRNA expression in bull sperm is correlated with higher sire conception rates (Arangasamy et al., 2011). As sperm are translationally silent, the mechanism by which sperm Crisp2 mRNA could directly regulate sperm function is unknown. Similarly, and many years earlier, CRISP2 deletions were proposed as causal of infertility in men with chromosomal aberrations (Olesen et al., 2001). These included a case of an Albanian man who presented with infertility, characterized by severe oligoasthenoteratospermia and testicular atrophy along with thickened basement membranes, wherein the CRISP2 gene, and others, was deleted via a t(6;21)(p21.1;p13) translocation (Paoloni-Giacobino et al., 2000). Further the screening of a group of infertile men with asthenozoospermia and teratozoospermia identified 21 CRISP2 polymorphisms (Jamsai et al., 2008b). Of particular note, one polymorphism, C196R, removed one of the strictly conserved cysteine residues within the hinge domain, which was shown to disrupt binding to gametogenetin 1 (GGN1) (see below), suggesting that C196R could compromise CRISP2 function and that correct disulphide bonding is critical for function (Jamsai et al., 2008b, 2008a). In terms of mechanism, several additional CRISP2-binding partners have been identified although their place in the overall context of CRISP2 function is yet to be settled. Specifically, CRISP2 can bind to mitogen-activated protein kinase kinase kinase 11 (MAP3K11) in the acrosome region of the developing spermatid and epididymal spermatozoa, via the ICR domain (Gibbs et al., 2007). As indicated above, CRISP2 also binds, via the ICR domain, to the carboxyl-most terminal of 158 amino-acid of GGN1 (Jamsai et al., 2008a). GGN1 also first appears in late pachytene spermatocytes and co-localizes with CRISP2 to sperm tail (Jamsai et al., 2008a). GGN1 also interacts with the Fanconi anaemia complex in spermatocytes where it is proposed to have a role in DNA repair (Jamsai et al., 2013), but the role of GGN1 and the significance of its interaction with CRISP2 in the sperm tail remains to be determined. A third binding partner of CRISP2 is the sperm head and tail-associated protein (SHTAP), which is incorporated into the head and tail of elongated spermatids and spermatozoa (Jamsai et al., 2010). The binding of CRISP2 to SHTAP is predominantly mediated by the CAP domain and stabilized by the hinge domain interaction (Gibbs et al., 2007; Jamsai et al., 2008a, 2010). The role of CAPs in epididymal maturation and post-ejaculate sperm function During epididymal transit, sperm come in to contact with CRISPs, which have been shown to play roles in the establishment of sperm functional competence in the female reproductive tract. These processes are collectively referred to as ‘epididymal sperm maturation’. Within the epididymis, CRISPs are produced by the principal cells and are found in high concentrations within epididymal fluid. CRISP1 is estimated to be at a concentration of ~12.4 μg/mg of total protein in the epididymis. Similarly, CRISP4 occurs at a concentration of ~18 μg/mg in the initial segment of the epididymis of mouse and 15.5 μg/mg in the rat (Eberspaecher et al., 1995; Krätzschmar et al., 1996). During epididymal transit, sperm bind CRISPs and this transfer is potentially mediated by epididymosomes. Epididymosomes are apocrine-generated extracellular microvesicles that modulate transfer of epididymal proteins onto the sperm (Martin-DeLeon, 2015; Sullivan, 2015). As mentioned above, the binding of CRISP to sperm occurs with two different affinities such that rat CRISP4 (published as CRISP1-D isoform) is loosely associated on the sperm surface and has been proposed as a decapacitation factor (Roberts et al., 2008). By contrast, the E isoform binds with high affinity to the sperm head and has been implicated in sperm–egg fusion (Roberts et al., 2003, 2008; Nixon et al., 2006; Cohen et al., 2007). Data indicate that CRISPs are required for optimal sperm function. Sperm from Crisp1−/− knockout mice have a compromised ability to manifest the acrosome reaction in response to progesterone stimulation and to penetrate the zona pellucida of intact oocytes. They also have reduced sperm progressive motility (Da Ros et al., 2008; Hu et al., 2018). Deletion of the Crisp4 gene in mice also results in altered sperm progressive motility, but interestingly the effect was to increase sperm velocity. These data indicate that CRISP1 and CRISP4 regulate different aspects of sperm motility. Similar to Crisp1−/− and Crisp2−/− knockout mice, sperm from Crisp4−/− mice also have a compromised ability to undergo the acrosome reaction in response to progesterone (Hu et al., 2018). In addition, Crisp4−/− knockout sperm had a reduced ability to undergo capacitation-associated tyrosine phosphorylation in a manner analogous to that seen in the Fer knockout mouse line, thus suggesting that these two proteins may function in the same pathway (Alvau et al., 2016; Hu et al., 2018). In contrast to the situation in mice, humans produce a single epididymal CRISP, namely CRISP1. As such, a better prediction of the role of human epididymal CRISPs may be provided through the analysis of Crisp1/4 double knockout mice i.e. total epididymal CRISP removal. Consistent with an upstream role for CRISP1 in regulating sperm motility, Crisp1−/− and Crisp1/4−/− double knockout mice both produced sperm with a significantly reduced ability for progressive motility (Hu et al., 2018). The removal of all epididymal CRISPs in mice also revealed a role for CRISPs in maintaining an immune-tolerant environment for the highly immunogenic sperm (Hu et al., 2018). The findings of this study were confirmed in a second study, which also reported defective epithelium function and an abnormal luminal pH (Carvajal et al., 2018). The loss of immunotolerance to sperm in Crisp1/4 double knockout mice bears a striking similarity to the role of the bacterial SCP domain-containing proteins in the induction of immunotolerance towards parasites, thus suggesting the CAP domain is responsible for this function. In terms of mechanism, rat CRISP4 (although incorrectly designated CRISP1 in the publication) can regulate calcium flow through the mouse CatSper channel (Ernesto et al., 2015). Consistent with the mouse data, rat CRISP4 can antagonize transient receptor potential (TRP) M8 ion channel function, where TRMP8 is localized on the sperm head and tail (Gibbs et al., 2011; Martínez-López et al., 2011) (discussed in detail below). Collectively, these data support a role of CRISP4 in regulating optimal sperm motility via calcium and potentially the phosphorylation of sperm tail proteins (Gibbs et al., 2011; Ernesto et al., 2015; Hu et al., 2018). The role of male-derived CAP proteins in the female reproductive tract Ejaculated sperm possess surface-associated components known as decapacitation factors, which suppress premature activation (capacitation) of the sperm in a reversible manner (Fraser, 1984; Fraser et al., 1990). Decapacitation factors include sterol components in the sperm plasma membrane and likely several associated proteins (Begley and Quinn, 1982; Bailey, 2010; Aitken and Nixon, 2013). A number of putative decapacitation factors has been characterized (Davis et al., 1980; Aitken and Nixon, 2013), including the glycosylated form of CRISP4 in rats (previously reported in literature as CRISP1) and potentially mouse CRISP1 (Nixon et al., 2006). The incubation of rat sperm with exogenous CRISP4 was shown to inhibit sperm tyrosine phosphorylation, capacitation and the ability to undergo the acrosome reaction in a dose-dependent manner (Roberts et al., 2003). As indicated above, data from Crisp4 knockout mice support CRISP4 being a primary regulator of capacitation and the downstream global tyrosine phosphorylation of sperm proteins (Hu et al., 2018); however, the data in these studies appear at direct odds with each other. Whether this difference is the result of using wild-type rat sperm compared to genetically modified (Crisp4 null) mouse sperm is not known. Although the exact mechanism is unknown, it has been suggested that CRISP4 may function as a ligand to a receptor on the sperm surface and that this pathway intersects with the activation of Fer kinase (Roberts et al., 2008; Alvau et al., 2016; Hu et al., 2018). In addition to the various roles for CRISPs on sperm function, evidence exists to suggest seminal plasma CRISPs, derived from the accessory glands, also play a role in optimizing male fertility. In mammals, seminal plasma is a fluid produced largely from the prostate and the seminal vesicles, although elements of testicular and epididymal products exist. It should be noted, however, that seminal plasma, as it is referred to clinically, may not reflect the in vivo situation wherein products from the different regions of the male reproductive tract are deposited into the female tract progressively [reviewed in (Ricardo, 2018)]. Elements of seminal plasma are critical for sperm transport, but also optimal sperm function and survival in the hostile environment of the female reproductive tract (Bromfield, 2014). Accumulating data, largely from horse, support a model whereby seminal plasma CRISPs play a crucial role in promoting male fertility in the female reproductive tract. CRISP3 is a major component of equine seminal plasma (1 mg/ml) and is produced in the seminal vesicles and the ampulla of the vas deferens of the mature male (Schambony et al., 1998; Giesecke et al., 2010; Gottschalk et al., 2016). CRISP3 SNPs were identified in Hanoverian warmblood horses and were found to predict a 7% decrease in pregnancy rate per cycle (Hamann et al., 2007; Usuga et al., 2018; Restrepo et al., 2019). These results were consistent and expanded in Colombian Creole horses (Hamann et al., 2007; Usuga et al., 2018; Restrepo et al., 2019). Collectively, these studies found a significant association between CRISP3 concentration in the ejaculate, or particular Crisp3 genotypes, and improved sperm parameters including vitality, membrane integrity and post-cryopreservation function. While the precise mechanism under-pinning these improvements is unknown, a study from Doty and colleagues identified CRISP3 as a suppressor of polymorphonuclear neutrophil binding to sperm (Doty et al., 2011). These data thus suggest that seminal plasma CRISP3 protects sperm from precocious destruction in the female reproductive tract following mating and prevent against excessive endometrial inflammatory responses to sperm antigens. Such a response is known as persistent mating-induced endometritis and is particularly common in horses and a significant cause of infertility (Christoffersen and Troedsson, 2017). A similar relationship between seminal plasma CRISP3 and fertility was observed in the camel (Waheed et al., 2015). In regard to the specific SNPs, two were associated with decreased fertility: E208K and Q239R (Hamann et al., 2007; Doty et al., 2011). Both reside in the ICR domain, and it is thus tempting to speculate that horse CRISP3 regulates as yet unidentified ion channels. The role of CAPs in the events of fertilization For successful sperm–egg fusion, the sperm undergoes re-organization of molecular complexes at the equatorial segment of the head followed by the acrosome reaction to gain fusion competence (Bedford and Rifkin, 1979; Inoue et al., 2005; Jones et al., 2008; Phopin et al., 2012; Robertson and Moldenhauer, 2014). It has been proposed that CRISP2, CRISP1 and/or GLIPR1L1 may be involved in this process. This hypothesis arose based on data from Busso et al. (Busso et al., 2005) showing that the addition of a peptide sequence corresponding to the CAP2 motif of CRISP2 interfered with sperm–egg fusion when added to IVF media (Busso et al., 2005). While these data are compatible with those from Maeda et al. showing that the N-terminal 101 amino acids of rat CRISP2 (including the CAP2 motif) can promote cell adhesion, no clarity is provided on the identity of the individual protein involved in vivo i.e. the CAP2 motif is common to all CAPs. Based on current knowledge, any/all of CRISP1, CRISP2 and GLIPR1L1 are plausible candidates to take part in this process. While each of the knockout mouse models for these genes have compromised fertility, upstream defects in acrosome function (and potentially capacitation) preclude a conclusive determination of a role at the site of fertilization (Gibbs et al., 2010; Hu et al., 2018; Lim et al., 2019). Similar results were obtained for rat CRISP4 (previously called CRISP1) using mouse and rat gametes (Cohen et al., 2000; Cohen et al., 2001, 2007; Maldera et al., 2014). However, the rat results should be cautiously interpreted as the study used mis-folded recombinant CRISP4 on mouse sperm. Controversy aside, CRISP2 is localized to the equatorial segment of sperm, which is the site of sperm-oocyte fusion (O’Bryan et al., 1998, 2001). A study of human sperm by Nimlamool and colleagues proposed that some of the CRISP2 released during acrosomal exocytosis re-associated with sperm at the equatorial segment, where it promotes sperm–egg fusion (Nimlamool et al., 2013). In addition, accumulating data from multiple species indicate that the GLIPR subfamily member, GLIPR1L1, plays a role in fertilization. GLIPR1L1 was first proposed to have a potential role in fertilization in mice when incubation with a GLIPR1L1 antibody was shown to significantly inhibit the binding of sperm to the zona pellucida in vitro (Gibbs et al., 2010). Concordant data were obtained in the bull (Caballero et al., 2012). Subsequently, GLIPR1L1 was identified as part of a multimeric protein complex associated with the bona fide sperm oocyte receptor protein, IZUMO1, in the mouse. Glipr1l1 gene deletion led to male subfertility characterized by sperm with a compromised ability to undergo the acrosome reaction and a failure of IZUMO1 relocalization during the acrosome reaction (Gibbs et al., 2010; Gaikwad et al., 2019). Collectively, these data suggest that GLIPR1L1 forms part of a multimeric protein complex, a likely association with lipid rafts, and that it plays a role in redistributing key proteins required for fertilization, including IZUMO1, during the final steps before oocyte binding. While GLIPR1L1 is not absolutely required for fertility, it appears to enhance the efficiency of fertilization. As mentioned above, GLIPR1L1 is also present on the plasma membrane of rat, bull and macaque sperm (Yudin et al., 2002; Caballero et al., 2012). It is interesting to note that the means by which GLIPR1L1 is associated with sperm differs between species. In the bull and macaque, GLIPR1L1 is produced in the epididymis and associated with lipid raft membrane domains via C-terminal GPI linkages (Caballero et al., 2012), whereas in mouse sperm, GLIPR1L1 is acquired during spermatogenesis and does not harbour a C-terminal GPI (Ren et al., 2006). In summary, these data suggest that at least one CAP protein can participate in the actual events of fertilization. As each of Crisp1, Crisp2, Crisp4 and Glipr1l1 knockout mice can achieve fertilization, these roles appear to be facilitative rather than essential (Gibbs et al., 2010; Hu et al., 2018; Gaikwad et al., 2019; Lim et al., 2019). The role of CAPs in ion channel regulation and sperm function Mature mammalian sperm contain many ion channels (Darszon et al., 1999; Acevedo et al., 2006). One of the most important is the sperm-specific, Ca2+-ion permeable, pH-dependent and low voltage-dependent ion channel known as CatSper (Ren et al., 2001; Lishko and Kirichok, 2010; Kirichok and Lishko, 2011; Sun et al., 2017) and its dysfunction leads to male sterility (Ren et al., 2001; Ren and Xia, 2010; Miller et al., 2015; Chung et al., 2017). CatSper deficient sperm are unable to manifest hyperactivated motility or penetrate the zona pellucida (Quill et al., 2003; Qi et al., 2007). In addition, they have slow, sluggish and defective undulatory movements when compared to the vigorous flagellar beating in wild-type sperm (Ren et al., 2001; Ren and Xia, 2010). Published data indicate that both rat CRISP4 (previously named CRISP1) and mouse CRISP2 can bind to CATSPER subunits (Lim et al., 2019). Consistent with this interpretation, sperm from Crisp2−/− mice have a stiff mid-piece and decreased progressive motility compared to sperm from wild-type littermates (Lim et al., 2019). In addition, CRISP2 can regulate Ca2+ flow through RyR, such that the CRISP domain activated both RyR1 and RyR2 at negative bilayer potentials and showed no effect at positive bilayer potentials when added to the luminal domain of the receptor. Mouse and rat CRISP4 were also shown to inhibit Ca2+ flow through the TRP ion channel TRPM8 (Gibbs et al., 2006, 2011; Martínez-López et al., 2011). As yet, the potential for CRISP1 or CRISP3 to regulate ion channel function on sperm has not been specifically assessed. Based on the range of ion channel types regulated by reptile CRISPs (Table I); however, we hypothesis that mammalian CRISPs will regulate many as yet undiscovered ion channel types. Table I Members of the CAP superfamily proteins and their proposed roles in reproduction and various physiological processes. CAP subfamilies . CAP subfamily protein . Species . Common name . Functional significance . References . Cysteine-rich secretory proteins CRISP1 Homo sapiens, Mus musculus, Rattus norvegicus Human, mouse, rat Regulates sperm capacitation, progressive motility, acrosome reaction and IVF. Regulates epididymal immune environment. Carvajal et al., 2018, Hu et al., 2018 CRISP2 Homo sapiens, Mus musculus, Rattus norvegicus Human, Mouse, Rat Regulates sperm progressive motility and acrosome reaction, optimal sperm flagellar beating, binds to CatSper subunit, regulates Ryanodine receptors. O’Bryan et al., 1998, O’Bryan et al., 2001, Lim et al., 2019 CRISP3 Homo sapiens, Mus musculus, Rattus norvegicus, Equus caballus, Camelus dromedarius Human, mouse, rat, horse, camel Potentially facilitates optimal mammalian fertilization. Upregulated in cancerous tissues, potential biomarker for autoimmune diseases Bjartell et al., 2007, Laine et al., 2007, Hamann et al., 2007, Ribeiro et al., 2011, Doty et al., 2011, Leng et al., 2018 CRISP4 Mus musculus, Rattus norvegicus Mouse, rat Regulates sperm capacitation, progressive motility, acrosome reaction and IVF, regulates epididymal immune environment, regulates CatSper and TRPM8 ion channel Cohen et al., 2000, Cohen et al., 2001, Gibbs et al., 2007, Carvajal et al., 2018, Hu et al., 2018 Helothermine Heloderma horridum Mexican beaded lizard Modulates voltage-gated calcium channels, voltage-gated potassium channels and ryanodine receptors Nobile et al., 1994, Morrissette et al., 1995, Nobile et al., 1996, CRVP Trimeresurus mucrosquamatus Brown spotted pit viper * Chang et al., 2013 Pseudechetoxin Pseudechis australis Mulga snake Blocks cyclic nucleotide-gated ion channels Brown et al., 1999, Suzuki et al., 2008 HG26 Hydrodynastes gigas Water cobra * Hill and Mackessy, 2000 Tigrin Rhabdophis tigrinus Olive-drab green snake * Yamazaki et al., 2003 Albomin Agkistrodon blomhoffi Japanese pit viper Blocks smooth muscle contraction Pseudecin Pseudechis porphyriacus Red-bellied black snake Blocks cyclic nucleotide-gated ion channels Yamazaki et al., 2003, Suzuki et al., 2008 Piscivorin Agkistrodon p. piscivorus Cottonmouth snake * Yamazaki et al., 2003 Ophanin Ophiophagus hannah King cobra * Catrin-1 Crotalus atrox Western diamondback rattlesnake * Catrin-2 * Cysteine-rich secretory proteins TJ-CRVP Trimeresurus jerdonii Jerdon’s pit viper * Jin et al., 2003 NA-CRVP1 Naja atra Chinese cobra * NA-CRVP2 * Latisemin Laticauda semifasciata Black-banded sea krait Inhibits cyclic nucleotide-gated ion channels Yamazaki and Morita, 2004 Trimucin Trimeresurus mucrosquamatus Brown spotted pit viper * Natrin Naja atra Chinese cobra Regulates variety of ion channels including Ca+ and K+ ion channels, Induces intercellular adhesion molecule-1 (ICAM-1), vascular adhesion molecule-1 (VCAM-1), and E-selectin expressions in ECs through the Zinc and HS modulations in mitogen-activated protein kinases (MAPKs) and nuclear factor (NF)-kB pathways. Wang et al., 2005, Wang et al., 2006, Wang et al., 2010 Stecrisp Trimeresurus stejnegeri Green pit viper Cysteine-rich domain with a K+ channel inhibitor-like fold Wang et al., 2005, Guo et al., 2005 Triflin Trimeresurus flavoviridi Kume Shima habu * Shikamoto et al., 2005 Patagonin Philodryas patagoniensis Green snake Skeletal myotoxic activity Peichoto et al., 2009 Crovirin Crotalus viridis Prairie rattlesnake Anti-protozoal activity Adade et al., 2014 ES-CRISP Echis carinatus sochureki Eastern saw-scaled viper negative regulator of the angiogenesis Lecht et al., 2015 Bj-CRP Bothrops jararaca Jararaca (large snake) Regulates inflammatory responses Lodovicho et al., 2016 Glioma pathogenesis related-1 proteins GLIPR1 Homo sapiens, Mus musculus Human, mouse Upregulated expression in cancerous tissues, P53-regulated gene and proapoptotic activity. Murphy et al., 1995, Rich et al., 1996, Chilukamarri et al., 2007 GLIPR1L1 Mus musculus, Bos taurus, Mouse, bull, monkey Regulates sperm acrosome reaction, associates with IZUMO to facilitate optimal sperm–egg binding, upregulated in cancerous tissues Yudin et al., 2002, Chilukamarri et al., 2007, Gibbs et al., 2010, Caballero et al., 2012, Gaikwad et al., 2019 GLIPR1L2 Homo sapiens, Mus musculus Human, mouse Expressed in diseased human tissues including prostate, bladder, kidney and bone marrow Ren et al., 2006 GLIPR1L3 Mus musculus Mouse * Ren et al., 2006 The Golgi-associated pathogenesis-related proteins GAPR Homo sapiens, Mus musculus Human, mouse Upregulated in mouse kidney fibrosis, immune regulation, amyloid regulation, sterol binding properties Kalluri et al., 2003, Desmoulière et al., 2005, van der Meer-Janssen et al., 2010, Shoji-Kawata et al., 2013 The Cysteine-Rich secretory protein LCCL domain-containing-1 proteins CRISPLD1 Homo sapiens, Mus musculus Human, mouse Associated with cardiovascular diseases Wilson et al., 2016, Wang et al., 2018 The Cysteine-Rich secretory protein LCCL domain-containing-2 proteins CRISPLD2 Homo sapiens, Mus musculus Human, mouse Associated with kidney and lung development, craniofacial morphogenesis, chronic bronchopulmonary dysplasia Nadeau et al., 2006, Quinlan et al., 2007, Lan et al., 2009, Shi et al., 2010, Mijiti et al., 2015 Peptidase inhibitor 15 proteins PI15 Homo sapiens, Mus musculus, Gallus Gallus domesticus Human, mouse, chicken Upregulated in dieseased kidney, associated with craniofacial development, anti-inflammatory and protease-like activity Takemoto et al., 2006, Falak et al., 2014, Nimmagadda et al., 2015, Prusty et al., 2018 Peptidase inhibitor 16 proteins PI16 Homo sapiens, Mus musculus Human, mouse Upregulated in cardiac diseased mouse model, associated with heart disease, inflammation Frost and Engelhardt, 2007, Rodríguez-Penas et al., 2015, Zhang et al., 2016, Regn et al., 2016, Hope et al., 2019 The C-type lectin proteins CLEC18A Homo sapiens, Mus musculus, Caenorhabditis elegans Human, mouse, C.elegans Immune response regulation, inflammation and autoimmunity, biomarker for chronic liver diseases, sterol binding properties Maeda et al., 1998, Miltsch et al., 2014, Huang et al., 2015, Liao et al., 2018 CLEC18B The venom allergen proteins Dol m 5 Dolichovespula maculate White-faced hornet Antigenic properties, suppresses host immune response, induces allergic response King et al., 1978, King et al., 1983, Li et al., 2001, Kovalick and Griffin, 2005, Justo Jacomini et al., 2014 Vesp c 5 Vespa crabo European hornet Pol a 5 Polistes annularis Paper wasp Ves v 5 Vespula vulgaris Yellow jacket Sol I 3 Solenopsis invicta Fire ants The bacterial SCP domain-containing proteins VAL Heligmosomoides polygyrus Intestinal roundworm Significantly upregulated during the parasitic phases. Immunomodulatory components Sotillo et al., 2014, Hunt et al., 2016 Ac-ASP-1 Ancylostoma caninum Dog hookworm Sterol binding activity, inhibits adhesion of activated human neutrophils to endothelial cells in the host and release of hydrogen peroxide from activated neutrophils Moyle et al., 1994, Cantacessi et al., 2009, Cantacessi and Gasser, 2012 Ac-ASP-2 NIF hg-VAP-1 Heterodera glycines Soybean cyst nematode Role in the infection of host plants Gao et al., 2001 The bacterial SCP domain-containing proteins hg-VAP-2 Mi-MSP-1 Meloidogyne incognita Cotton root-knot nematode Role in the infection of host plants Ding et al., 2000 Mi-VAP-2 HpVAL-4 Heligmosomoides polygyrus bakeri Parasitic hookworm Possess a caveolin-binding motif (CBM) and palmitate binding cavity, which has the ability to bind to cholesterol and fatty acid Ma et al., 2011 SmVAL1-SmVAL13 Schistosoma mansoni Water-borne parasite Role in stress responses. Protects the parasite against hosts defense mechanisms Basch and DiConza, 1974, Chalmers et al., 2008, Wu et al., 2009, Cantacessi and Gasser, 2012 Ov-ASP-1 Onchocerca volvulus Onchocerca Induce angiogenesis, supports parasite maturation during parasitism Tawe et al., 2000, Lizotte-Waniewski et al., 2000 Ov-ASP-2 Ov-ASP-3 The pathogenesis-related-1 proteins Pr-1a Nicotiana tabacum, Arabidopsis thaliana, Helianthus annuus, Pisum sativum Tobacco, Arabidopsis, sunflower, peas Role in the surveillance mechanism in plant immune system, sterol-binding properties van Loon, 1985, Elvira et al., 2008, Chaki et al., 2009, Begara-Morales et al., 2013, Takahashi et al., 2016 The fungal pathogenesis-related proteins PRY1 Saccharomyces cerevisiae S. cerevisiae Sterol and fatty acid-binding properties Choudhary and Schneiter, 2012, Choudhary et al., 2014 PRY2 CAP subfamilies . CAP subfamily protein . Species . Common name . Functional significance . References . Cysteine-rich secretory proteins CRISP1 Homo sapiens, Mus musculus, Rattus norvegicus Human, mouse, rat Regulates sperm capacitation, progressive motility, acrosome reaction and IVF. Regulates epididymal immune environment. Carvajal et al., 2018, Hu et al., 2018 CRISP2 Homo sapiens, Mus musculus, Rattus norvegicus Human, Mouse, Rat Regulates sperm progressive motility and acrosome reaction, optimal sperm flagellar beating, binds to CatSper subunit, regulates Ryanodine receptors. O’Bryan et al., 1998, O’Bryan et al., 2001, Lim et al., 2019 CRISP3 Homo sapiens, Mus musculus, Rattus norvegicus, Equus caballus, Camelus dromedarius Human, mouse, rat, horse, camel Potentially facilitates optimal mammalian fertilization. Upregulated in cancerous tissues, potential biomarker for autoimmune diseases Bjartell et al., 2007, Laine et al., 2007, Hamann et al., 2007, Ribeiro et al., 2011, Doty et al., 2011, Leng et al., 2018 CRISP4 Mus musculus, Rattus norvegicus Mouse, rat Regulates sperm capacitation, progressive motility, acrosome reaction and IVF, regulates epididymal immune environment, regulates CatSper and TRPM8 ion channel Cohen et al., 2000, Cohen et al., 2001, Gibbs et al., 2007, Carvajal et al., 2018, Hu et al., 2018 Helothermine Heloderma horridum Mexican beaded lizard Modulates voltage-gated calcium channels, voltage-gated potassium channels and ryanodine receptors Nobile et al., 1994, Morrissette et al., 1995, Nobile et al., 1996, CRVP Trimeresurus mucrosquamatus Brown spotted pit viper * Chang et al., 2013 Pseudechetoxin Pseudechis australis Mulga snake Blocks cyclic nucleotide-gated ion channels Brown et al., 1999, Suzuki et al., 2008 HG26 Hydrodynastes gigas Water cobra * Hill and Mackessy, 2000 Tigrin Rhabdophis tigrinus Olive-drab green snake * Yamazaki et al., 2003 Albomin Agkistrodon blomhoffi Japanese pit viper Blocks smooth muscle contraction Pseudecin Pseudechis porphyriacus Red-bellied black snake Blocks cyclic nucleotide-gated ion channels Yamazaki et al., 2003, Suzuki et al., 2008 Piscivorin Agkistrodon p. piscivorus Cottonmouth snake * Yamazaki et al., 2003 Ophanin Ophiophagus hannah King cobra * Catrin-1 Crotalus atrox Western diamondback rattlesnake * Catrin-2 * Cysteine-rich secretory proteins TJ-CRVP Trimeresurus jerdonii Jerdon’s pit viper * Jin et al., 2003 NA-CRVP1 Naja atra Chinese cobra * NA-CRVP2 * Latisemin Laticauda semifasciata Black-banded sea krait Inhibits cyclic nucleotide-gated ion channels Yamazaki and Morita, 2004 Trimucin Trimeresurus mucrosquamatus Brown spotted pit viper * Natrin Naja atra Chinese cobra Regulates variety of ion channels including Ca+ and K+ ion channels, Induces intercellular adhesion molecule-1 (ICAM-1), vascular adhesion molecule-1 (VCAM-1), and E-selectin expressions in ECs through the Zinc and HS modulations in mitogen-activated protein kinases (MAPKs) and nuclear factor (NF)-kB pathways. Wang et al., 2005, Wang et al., 2006, Wang et al., 2010 Stecrisp Trimeresurus stejnegeri Green pit viper Cysteine-rich domain with a K+ channel inhibitor-like fold Wang et al., 2005, Guo et al., 2005 Triflin Trimeresurus flavoviridi Kume Shima habu * Shikamoto et al., 2005 Patagonin Philodryas patagoniensis Green snake Skeletal myotoxic activity Peichoto et al., 2009 Crovirin Crotalus viridis Prairie rattlesnake Anti-protozoal activity Adade et al., 2014 ES-CRISP Echis carinatus sochureki Eastern saw-scaled viper negative regulator of the angiogenesis Lecht et al., 2015 Bj-CRP Bothrops jararaca Jararaca (large snake) Regulates inflammatory responses Lodovicho et al., 2016 Glioma pathogenesis related-1 proteins GLIPR1 Homo sapiens, Mus musculus Human, mouse Upregulated expression in cancerous tissues, P53-regulated gene and proapoptotic activity. Murphy et al., 1995, Rich et al., 1996, Chilukamarri et al., 2007 GLIPR1L1 Mus musculus, Bos taurus, Mouse, bull, monkey Regulates sperm acrosome reaction, associates with IZUMO to facilitate optimal sperm–egg binding, upregulated in cancerous tissues Yudin et al., 2002, Chilukamarri et al., 2007, Gibbs et al., 2010, Caballero et al., 2012, Gaikwad et al., 2019 GLIPR1L2 Homo sapiens, Mus musculus Human, mouse Expressed in diseased human tissues including prostate, bladder, kidney and bone marrow Ren et al., 2006 GLIPR1L3 Mus musculus Mouse * Ren et al., 2006 The Golgi-associated pathogenesis-related proteins GAPR Homo sapiens, Mus musculus Human, mouse Upregulated in mouse kidney fibrosis, immune regulation, amyloid regulation, sterol binding properties Kalluri et al., 2003, Desmoulière et al., 2005, van der Meer-Janssen et al., 2010, Shoji-Kawata et al., 2013 The Cysteine-Rich secretory protein LCCL domain-containing-1 proteins CRISPLD1 Homo sapiens, Mus musculus Human, mouse Associated with cardiovascular diseases Wilson et al., 2016, Wang et al., 2018 The Cysteine-Rich secretory protein LCCL domain-containing-2 proteins CRISPLD2 Homo sapiens, Mus musculus Human, mouse Associated with kidney and lung development, craniofacial morphogenesis, chronic bronchopulmonary dysplasia Nadeau et al., 2006, Quinlan et al., 2007, Lan et al., 2009, Shi et al., 2010, Mijiti et al., 2015 Peptidase inhibitor 15 proteins PI15 Homo sapiens, Mus musculus, Gallus Gallus domesticus Human, mouse, chicken Upregulated in dieseased kidney, associated with craniofacial development, anti-inflammatory and protease-like activity Takemoto et al., 2006, Falak et al., 2014, Nimmagadda et al., 2015, Prusty et al., 2018 Peptidase inhibitor 16 proteins PI16 Homo sapiens, Mus musculus Human, mouse Upregulated in cardiac diseased mouse model, associated with heart disease, inflammation Frost and Engelhardt, 2007, Rodríguez-Penas et al., 2015, Zhang et al., 2016, Regn et al., 2016, Hope et al., 2019 The C-type lectin proteins CLEC18A Homo sapiens, Mus musculus, Caenorhabditis elegans Human, mouse, C.elegans Immune response regulation, inflammation and autoimmunity, biomarker for chronic liver diseases, sterol binding properties Maeda et al., 1998, Miltsch et al., 2014, Huang et al., 2015, Liao et al., 2018 CLEC18B The venom allergen proteins Dol m 5 Dolichovespula maculate White-faced hornet Antigenic properties, suppresses host immune response, induces allergic response King et al., 1978, King et al., 1983, Li et al., 2001, Kovalick and Griffin, 2005, Justo Jacomini et al., 2014 Vesp c 5 Vespa crabo European hornet Pol a 5 Polistes annularis Paper wasp Ves v 5 Vespula vulgaris Yellow jacket Sol I 3 Solenopsis invicta Fire ants The bacterial SCP domain-containing proteins VAL Heligmosomoides polygyrus Intestinal roundworm Significantly upregulated during the parasitic phases. Immunomodulatory components Sotillo et al., 2014, Hunt et al., 2016 Ac-ASP-1 Ancylostoma caninum Dog hookworm Sterol binding activity, inhibits adhesion of activated human neutrophils to endothelial cells in the host and release of hydrogen peroxide from activated neutrophils Moyle et al., 1994, Cantacessi et al., 2009, Cantacessi and Gasser, 2012 Ac-ASP-2 NIF hg-VAP-1 Heterodera glycines Soybean cyst nematode Role in the infection of host plants Gao et al., 2001 The bacterial SCP domain-containing proteins hg-VAP-2 Mi-MSP-1 Meloidogyne incognita Cotton root-knot nematode Role in the infection of host plants Ding et al., 2000 Mi-VAP-2 HpVAL-4 Heligmosomoides polygyrus bakeri Parasitic hookworm Possess a caveolin-binding motif (CBM) and palmitate binding cavity, which has the ability to bind to cholesterol and fatty acid Ma et al., 2011 SmVAL1-SmVAL13 Schistosoma mansoni Water-borne parasite Role in stress responses. Protects the parasite against hosts defense mechanisms Basch and DiConza, 1974, Chalmers et al., 2008, Wu et al., 2009, Cantacessi and Gasser, 2012 Ov-ASP-1 Onchocerca volvulus Onchocerca Induce angiogenesis, supports parasite maturation during parasitism Tawe et al., 2000, Lizotte-Waniewski et al., 2000 Ov-ASP-2 Ov-ASP-3 The pathogenesis-related-1 proteins Pr-1a Nicotiana tabacum, Arabidopsis thaliana, Helianthus annuus, Pisum sativum Tobacco, Arabidopsis, sunflower, peas Role in the surveillance mechanism in plant immune system, sterol-binding properties van Loon, 1985, Elvira et al., 2008, Chaki et al., 2009, Begara-Morales et al., 2013, Takahashi et al., 2016 The fungal pathogenesis-related proteins PRY1 Saccharomyces cerevisiae S. cerevisiae Sterol and fatty acid-binding properties Choudhary and Schneiter, 2012, Choudhary et al., 2014 PRY2 *Indicates unknown. ASP: anclystoma-secreted proteins; CAP: cysteine-rich secretory proteins, antigen 5, and pathogenesis-related 1; CatSper: cation channels of sperm; CLEC: C-type lectin; CRISP: cysteine-rich secretory proteins; CRISPLD: cysteine-rich LCCL domain-containing; CRVP: cysteine-rich venom protein; GAPR: Golgi-associated pathogenesis-related; GLIPR: glioma pathogenesis-related; GLIPR1L1: glioma pathogenesis-related 1 like 1; IVF: in vitro fertilization; NIF: neutrophil inhibitory factor; PI: peptidase inhibitor; Pr-1: pathogenesis-related 1; PRY: fungal pathogenesis-related; TRP: transient receptor potential; VAH: venom allergen-homologous; VAL: venom allergen-like; VAP: venom allergen-proteins Open in new tab Table I Members of the CAP superfamily proteins and their proposed roles in reproduction and various physiological processes. CAP subfamilies . CAP subfamily protein . Species . Common name . Functional significance . References . Cysteine-rich secretory proteins CRISP1 Homo sapiens, Mus musculus, Rattus norvegicus Human, mouse, rat Regulates sperm capacitation, progressive motility, acrosome reaction and IVF. Regulates epididymal immune environment. Carvajal et al., 2018, Hu et al., 2018 CRISP2 Homo sapiens, Mus musculus, Rattus norvegicus Human, Mouse, Rat Regulates sperm progressive motility and acrosome reaction, optimal sperm flagellar beating, binds to CatSper subunit, regulates Ryanodine receptors. O’Bryan et al., 1998, O’Bryan et al., 2001, Lim et al., 2019 CRISP3 Homo sapiens, Mus musculus, Rattus norvegicus, Equus caballus, Camelus dromedarius Human, mouse, rat, horse, camel Potentially facilitates optimal mammalian fertilization. Upregulated in cancerous tissues, potential biomarker for autoimmune diseases Bjartell et al., 2007, Laine et al., 2007, Hamann et al., 2007, Ribeiro et al., 2011, Doty et al., 2011, Leng et al., 2018 CRISP4 Mus musculus, Rattus norvegicus Mouse, rat Regulates sperm capacitation, progressive motility, acrosome reaction and IVF, regulates epididymal immune environment, regulates CatSper and TRPM8 ion channel Cohen et al., 2000, Cohen et al., 2001, Gibbs et al., 2007, Carvajal et al., 2018, Hu et al., 2018 Helothermine Heloderma horridum Mexican beaded lizard Modulates voltage-gated calcium channels, voltage-gated potassium channels and ryanodine receptors Nobile et al., 1994, Morrissette et al., 1995, Nobile et al., 1996, CRVP Trimeresurus mucrosquamatus Brown spotted pit viper * Chang et al., 2013 Pseudechetoxin Pseudechis australis Mulga snake Blocks cyclic nucleotide-gated ion channels Brown et al., 1999, Suzuki et al., 2008 HG26 Hydrodynastes gigas Water cobra * Hill and Mackessy, 2000 Tigrin Rhabdophis tigrinus Olive-drab green snake * Yamazaki et al., 2003 Albomin Agkistrodon blomhoffi Japanese pit viper Blocks smooth muscle contraction Pseudecin Pseudechis porphyriacus Red-bellied black snake Blocks cyclic nucleotide-gated ion channels Yamazaki et al., 2003, Suzuki et al., 2008 Piscivorin Agkistrodon p. piscivorus Cottonmouth snake * Yamazaki et al., 2003 Ophanin Ophiophagus hannah King cobra * Catrin-1 Crotalus atrox Western diamondback rattlesnake * Catrin-2 * Cysteine-rich secretory proteins TJ-CRVP Trimeresurus jerdonii Jerdon’s pit viper * Jin et al., 2003 NA-CRVP1 Naja atra Chinese cobra * NA-CRVP2 * Latisemin Laticauda semifasciata Black-banded sea krait Inhibits cyclic nucleotide-gated ion channels Yamazaki and Morita, 2004 Trimucin Trimeresurus mucrosquamatus Brown spotted pit viper * Natrin Naja atra Chinese cobra Regulates variety of ion channels including Ca+ and K+ ion channels, Induces intercellular adhesion molecule-1 (ICAM-1), vascular adhesion molecule-1 (VCAM-1), and E-selectin expressions in ECs through the Zinc and HS modulations in mitogen-activated protein kinases (MAPKs) and nuclear factor (NF)-kB pathways. Wang et al., 2005, Wang et al., 2006, Wang et al., 2010 Stecrisp Trimeresurus stejnegeri Green pit viper Cysteine-rich domain with a K+ channel inhibitor-like fold Wang et al., 2005, Guo et al., 2005 Triflin Trimeresurus flavoviridi Kume Shima habu * Shikamoto et al., 2005 Patagonin Philodryas patagoniensis Green snake Skeletal myotoxic activity Peichoto et al., 2009 Crovirin Crotalus viridis Prairie rattlesnake Anti-protozoal activity Adade et al., 2014 ES-CRISP Echis carinatus sochureki Eastern saw-scaled viper negative regulator of the angiogenesis Lecht et al., 2015 Bj-CRP Bothrops jararaca Jararaca (large snake) Regulates inflammatory responses Lodovicho et al., 2016 Glioma pathogenesis related-1 proteins GLIPR1 Homo sapiens, Mus musculus Human, mouse Upregulated expression in cancerous tissues, P53-regulated gene and proapoptotic activity. Murphy et al., 1995, Rich et al., 1996, Chilukamarri et al., 2007 GLIPR1L1 Mus musculus, Bos taurus, Mouse, bull, monkey Regulates sperm acrosome reaction, associates with IZUMO to facilitate optimal sperm–egg binding, upregulated in cancerous tissues Yudin et al., 2002, Chilukamarri et al., 2007, Gibbs et al., 2010, Caballero et al., 2012, Gaikwad et al., 2019 GLIPR1L2 Homo sapiens, Mus musculus Human, mouse Expressed in diseased human tissues including prostate, bladder, kidney and bone marrow Ren et al., 2006 GLIPR1L3 Mus musculus Mouse * Ren et al., 2006 The Golgi-associated pathogenesis-related proteins GAPR Homo sapiens, Mus musculus Human, mouse Upregulated in mouse kidney fibrosis, immune regulation, amyloid regulation, sterol binding properties Kalluri et al., 2003, Desmoulière et al., 2005, van der Meer-Janssen et al., 2010, Shoji-Kawata et al., 2013 The Cysteine-Rich secretory protein LCCL domain-containing-1 proteins CRISPLD1 Homo sapiens, Mus musculus Human, mouse Associated with cardiovascular diseases Wilson et al., 2016, Wang et al., 2018 The Cysteine-Rich secretory protein LCCL domain-containing-2 proteins CRISPLD2 Homo sapiens, Mus musculus Human, mouse Associated with kidney and lung development, craniofacial morphogenesis, chronic bronchopulmonary dysplasia Nadeau et al., 2006, Quinlan et al., 2007, Lan et al., 2009, Shi et al., 2010, Mijiti et al., 2015 Peptidase inhibitor 15 proteins PI15 Homo sapiens, Mus musculus, Gallus Gallus domesticus Human, mouse, chicken Upregulated in dieseased kidney, associated with craniofacial development, anti-inflammatory and protease-like activity Takemoto et al., 2006, Falak et al., 2014, Nimmagadda et al., 2015, Prusty et al., 2018 Peptidase inhibitor 16 proteins PI16 Homo sapiens, Mus musculus Human, mouse Upregulated in cardiac diseased mouse model, associated with heart disease, inflammation Frost and Engelhardt, 2007, Rodríguez-Penas et al., 2015, Zhang et al., 2016, Regn et al., 2016, Hope et al., 2019 The C-type lectin proteins CLEC18A Homo sapiens, Mus musculus, Caenorhabditis elegans Human, mouse, C.elegans Immune response regulation, inflammation and autoimmunity, biomarker for chronic liver diseases, sterol binding properties Maeda et al., 1998, Miltsch et al., 2014, Huang et al., 2015, Liao et al., 2018 CLEC18B The venom allergen proteins Dol m 5 Dolichovespula maculate White-faced hornet Antigenic properties, suppresses host immune response, induces allergic response King et al., 1978, King et al., 1983, Li et al., 2001, Kovalick and Griffin, 2005, Justo Jacomini et al., 2014 Vesp c 5 Vespa crabo European hornet Pol a 5 Polistes annularis Paper wasp Ves v 5 Vespula vulgaris Yellow jacket Sol I 3 Solenopsis invicta Fire ants The bacterial SCP domain-containing proteins VAL Heligmosomoides polygyrus Intestinal roundworm Significantly upregulated during the parasitic phases. Immunomodulatory components Sotillo et al., 2014, Hunt et al., 2016 Ac-ASP-1 Ancylostoma caninum Dog hookworm Sterol binding activity, inhibits adhesion of activated human neutrophils to endothelial cells in the host and release of hydrogen peroxide from activated neutrophils Moyle et al., 1994, Cantacessi et al., 2009, Cantacessi and Gasser, 2012 Ac-ASP-2 NIF hg-VAP-1 Heterodera glycines Soybean cyst nematode Role in the infection of host plants Gao et al., 2001 The bacterial SCP domain-containing proteins hg-VAP-2 Mi-MSP-1 Meloidogyne incognita Cotton root-knot nematode Role in the infection of host plants Ding et al., 2000 Mi-VAP-2 HpVAL-4 Heligmosomoides polygyrus bakeri Parasitic hookworm Possess a caveolin-binding motif (CBM) and palmitate binding cavity, which has the ability to bind to cholesterol and fatty acid Ma et al., 2011 SmVAL1-SmVAL13 Schistosoma mansoni Water-borne parasite Role in stress responses. Protects the parasite against hosts defense mechanisms Basch and DiConza, 1974, Chalmers et al., 2008, Wu et al., 2009, Cantacessi and Gasser, 2012 Ov-ASP-1 Onchocerca volvulus Onchocerca Induce angiogenesis, supports parasite maturation during parasitism Tawe et al., 2000, Lizotte-Waniewski et al., 2000 Ov-ASP-2 Ov-ASP-3 The pathogenesis-related-1 proteins Pr-1a Nicotiana tabacum, Arabidopsis thaliana, Helianthus annuus, Pisum sativum Tobacco, Arabidopsis, sunflower, peas Role in the surveillance mechanism in plant immune system, sterol-binding properties van Loon, 1985, Elvira et al., 2008, Chaki et al., 2009, Begara-Morales et al., 2013, Takahashi et al., 2016 The fungal pathogenesis-related proteins PRY1 Saccharomyces cerevisiae S. cerevisiae Sterol and fatty acid-binding properties Choudhary and Schneiter, 2012, Choudhary et al., 2014 PRY2 CAP subfamilies . CAP subfamily protein . Species . Common name . Functional significance . References . Cysteine-rich secretory proteins CRISP1 Homo sapiens, Mus musculus, Rattus norvegicus Human, mouse, rat Regulates sperm capacitation, progressive motility, acrosome reaction and IVF. Regulates epididymal immune environment. Carvajal et al., 2018, Hu et al., 2018 CRISP2 Homo sapiens, Mus musculus, Rattus norvegicus Human, Mouse, Rat Regulates sperm progressive motility and acrosome reaction, optimal sperm flagellar beating, binds to CatSper subunit, regulates Ryanodine receptors. O’Bryan et al., 1998, O’Bryan et al., 2001, Lim et al., 2019 CRISP3 Homo sapiens, Mus musculus, Rattus norvegicus, Equus caballus, Camelus dromedarius Human, mouse, rat, horse, camel Potentially facilitates optimal mammalian fertilization. Upregulated in cancerous tissues, potential biomarker for autoimmune diseases Bjartell et al., 2007, Laine et al., 2007, Hamann et al., 2007, Ribeiro et al., 2011, Doty et al., 2011, Leng et al., 2018 CRISP4 Mus musculus, Rattus norvegicus Mouse, rat Regulates sperm capacitation, progressive motility, acrosome reaction and IVF, regulates epididymal immune environment, regulates CatSper and TRPM8 ion channel Cohen et al., 2000, Cohen et al., 2001, Gibbs et al., 2007, Carvajal et al., 2018, Hu et al., 2018 Helothermine Heloderma horridum Mexican beaded lizard Modulates voltage-gated calcium channels, voltage-gated potassium channels and ryanodine receptors Nobile et al., 1994, Morrissette et al., 1995, Nobile et al., 1996, CRVP Trimeresurus mucrosquamatus Brown spotted pit viper * Chang et al., 2013 Pseudechetoxin Pseudechis australis Mulga snake Blocks cyclic nucleotide-gated ion channels Brown et al., 1999, Suzuki et al., 2008 HG26 Hydrodynastes gigas Water cobra * Hill and Mackessy, 2000 Tigrin Rhabdophis tigrinus Olive-drab green snake * Yamazaki et al., 2003 Albomin Agkistrodon blomhoffi Japanese pit viper Blocks smooth muscle contraction Pseudecin Pseudechis porphyriacus Red-bellied black snake Blocks cyclic nucleotide-gated ion channels Yamazaki et al., 2003, Suzuki et al., 2008 Piscivorin Agkistrodon p. piscivorus Cottonmouth snake * Yamazaki et al., 2003 Ophanin Ophiophagus hannah King cobra * Catrin-1 Crotalus atrox Western diamondback rattlesnake * Catrin-2 * Cysteine-rich secretory proteins TJ-CRVP Trimeresurus jerdonii Jerdon’s pit viper * Jin et al., 2003 NA-CRVP1 Naja atra Chinese cobra * NA-CRVP2 * Latisemin Laticauda semifasciata Black-banded sea krait Inhibits cyclic nucleotide-gated ion channels Yamazaki and Morita, 2004 Trimucin Trimeresurus mucrosquamatus Brown spotted pit viper * Natrin Naja atra Chinese cobra Regulates variety of ion channels including Ca+ and K+ ion channels, Induces intercellular adhesion molecule-1 (ICAM-1), vascular adhesion molecule-1 (VCAM-1), and E-selectin expressions in ECs through the Zinc and HS modulations in mitogen-activated protein kinases (MAPKs) and nuclear factor (NF)-kB pathways. Wang et al., 2005, Wang et al., 2006, Wang et al., 2010 Stecrisp Trimeresurus stejnegeri Green pit viper Cysteine-rich domain with a K+ channel inhibitor-like fold Wang et al., 2005, Guo et al., 2005 Triflin Trimeresurus flavoviridi Kume Shima habu * Shikamoto et al., 2005 Patagonin Philodryas patagoniensis Green snake Skeletal myotoxic activity Peichoto et al., 2009 Crovirin Crotalus viridis Prairie rattlesnake Anti-protozoal activity Adade et al., 2014 ES-CRISP Echis carinatus sochureki Eastern saw-scaled viper negative regulator of the angiogenesis Lecht et al., 2015 Bj-CRP Bothrops jararaca Jararaca (large snake) Regulates inflammatory responses Lodovicho et al., 2016 Glioma pathogenesis related-1 proteins GLIPR1 Homo sapiens, Mus musculus Human, mouse Upregulated expression in cancerous tissues, P53-regulated gene and proapoptotic activity. Murphy et al., 1995, Rich et al., 1996, Chilukamarri et al., 2007 GLIPR1L1 Mus musculus, Bos taurus, Mouse, bull, monkey Regulates sperm acrosome reaction, associates with IZUMO to facilitate optimal sperm–egg binding, upregulated in cancerous tissues Yudin et al., 2002, Chilukamarri et al., 2007, Gibbs et al., 2010, Caballero et al., 2012, Gaikwad et al., 2019 GLIPR1L2 Homo sapiens, Mus musculus Human, mouse Expressed in diseased human tissues including prostate, bladder, kidney and bone marrow Ren et al., 2006 GLIPR1L3 Mus musculus Mouse * Ren et al., 2006 The Golgi-associated pathogenesis-related proteins GAPR Homo sapiens, Mus musculus Human, mouse Upregulated in mouse kidney fibrosis, immune regulation, amyloid regulation, sterol binding properties Kalluri et al., 2003, Desmoulière et al., 2005, van der Meer-Janssen et al., 2010, Shoji-Kawata et al., 2013 The Cysteine-Rich secretory protein LCCL domain-containing-1 proteins CRISPLD1 Homo sapiens, Mus musculus Human, mouse Associated with cardiovascular diseases Wilson et al., 2016, Wang et al., 2018 The Cysteine-Rich secretory protein LCCL domain-containing-2 proteins CRISPLD2 Homo sapiens, Mus musculus Human, mouse Associated with kidney and lung development, craniofacial morphogenesis, chronic bronchopulmonary dysplasia Nadeau et al., 2006, Quinlan et al., 2007, Lan et al., 2009, Shi et al., 2010, Mijiti et al., 2015 Peptidase inhibitor 15 proteins PI15 Homo sapiens, Mus musculus, Gallus Gallus domesticus Human, mouse, chicken Upregulated in dieseased kidney, associated with craniofacial development, anti-inflammatory and protease-like activity Takemoto et al., 2006, Falak et al., 2014, Nimmagadda et al., 2015, Prusty et al., 2018 Peptidase inhibitor 16 proteins PI16 Homo sapiens, Mus musculus Human, mouse Upregulated in cardiac diseased mouse model, associated with heart disease, inflammation Frost and Engelhardt, 2007, Rodríguez-Penas et al., 2015, Zhang et al., 2016, Regn et al., 2016, Hope et al., 2019 The C-type lectin proteins CLEC18A Homo sapiens, Mus musculus, Caenorhabditis elegans Human, mouse, C.elegans Immune response regulation, inflammation and autoimmunity, biomarker for chronic liver diseases, sterol binding properties Maeda et al., 1998, Miltsch et al., 2014, Huang et al., 2015, Liao et al., 2018 CLEC18B The venom allergen proteins Dol m 5 Dolichovespula maculate White-faced hornet Antigenic properties, suppresses host immune response, induces allergic response King et al., 1978, King et al., 1983, Li et al., 2001, Kovalick and Griffin, 2005, Justo Jacomini et al., 2014 Vesp c 5 Vespa crabo European hornet Pol a 5 Polistes annularis Paper wasp Ves v 5 Vespula vulgaris Yellow jacket Sol I 3 Solenopsis invicta Fire ants The bacterial SCP domain-containing proteins VAL Heligmosomoides polygyrus Intestinal roundworm Significantly upregulated during the parasitic phases. Immunomodulatory components Sotillo et al., 2014, Hunt et al., 2016 Ac-ASP-1 Ancylostoma caninum Dog hookworm Sterol binding activity, inhibits adhesion of activated human neutrophils to endothelial cells in the host and release of hydrogen peroxide from activated neutrophils Moyle et al., 1994, Cantacessi et al., 2009, Cantacessi and Gasser, 2012 Ac-ASP-2 NIF hg-VAP-1 Heterodera glycines Soybean cyst nematode Role in the infection of host plants Gao et al., 2001 The bacterial SCP domain-containing proteins hg-VAP-2 Mi-MSP-1 Meloidogyne incognita Cotton root-knot nematode Role in the infection of host plants Ding et al., 2000 Mi-VAP-2 HpVAL-4 Heligmosomoides polygyrus bakeri Parasitic hookworm Possess a caveolin-binding motif (CBM) and palmitate binding cavity, which has the ability to bind to cholesterol and fatty acid Ma et al., 2011 SmVAL1-SmVAL13 Schistosoma mansoni Water-borne parasite Role in stress responses. Protects the parasite against hosts defense mechanisms Basch and DiConza, 1974, Chalmers et al., 2008, Wu et al., 2009, Cantacessi and Gasser, 2012 Ov-ASP-1 Onchocerca volvulus Onchocerca Induce angiogenesis, supports parasite maturation during parasitism Tawe et al., 2000, Lizotte-Waniewski et al., 2000 Ov-ASP-2 Ov-ASP-3 The pathogenesis-related-1 proteins Pr-1a Nicotiana tabacum, Arabidopsis thaliana, Helianthus annuus, Pisum sativum Tobacco, Arabidopsis, sunflower, peas Role in the surveillance mechanism in plant immune system, sterol-binding properties van Loon, 1985, Elvira et al., 2008, Chaki et al., 2009, Begara-Morales et al., 2013, Takahashi et al., 2016 The fungal pathogenesis-related proteins PRY1 Saccharomyces cerevisiae S. cerevisiae Sterol and fatty acid-binding properties Choudhary and Schneiter, 2012, Choudhary et al., 2014 PRY2 *Indicates unknown. ASP: anclystoma-secreted proteins; CAP: cysteine-rich secretory proteins, antigen 5, and pathogenesis-related 1; CatSper: cation channels of sperm; CLEC: C-type lectin; CRISP: cysteine-rich secretory proteins; CRISPLD: cysteine-rich LCCL domain-containing; CRVP: cysteine-rich venom protein; GAPR: Golgi-associated pathogenesis-related; GLIPR: glioma pathogenesis-related; GLIPR1L1: glioma pathogenesis-related 1 like 1; IVF: in vitro fertilization; NIF: neutrophil inhibitory factor; PI: peptidase inhibitor; Pr-1: pathogenesis-related 1; PRY: fungal pathogenesis-related; TRP: transient receptor potential; VAH: venom allergen-homologous; VAL: venom allergen-like; VAP: venom allergen-proteins Open in new tab The role of CAPs as a sperm chemoattractant Sperm chemotaxis is one of the early signalling events between sperm and the oocytes (Sawada et al., 2014). These signals are mediated by the proteins released from the female gametes and are thought to guide the sperm through the environment or female reproductive tract (Burnett et al., 2011). Sperm chemoattraction has been extensively studied in invertebrates, such as sea urchins, molluscs and corals, but is poorly characterized in mammals (Ishikawa et al., 2004; Krug et al., 2009; Morita et al., 2009; Chang et al., 2013). In many animal species, sperm chemotaxis is mediated by the extracellular matrix layer surrounding oocytes (Wassarman and Litscher, 2009; Ikawa et al., 2010) For example, the vitelline coat of ascidians plays a crucial role in mediating sperm–egg communication by guiding the sperm to the egg surface. Ascidians, or sea squirts, are hermaphroditic animals, which release both male and female gametes at the same time during spawning season (Fuke, 1983; Ben-Shlomo, 2008). However, in order to achieve genetic diversity, most ascidians exhibit self-sterility or allogeneic fertilization (Jiang and Smith, 2005). Numerous studies, spanning several species, indicate that CAP proteins play a role in this process. For example, in Halocynthia roretzi, the sea pineapple, studies have shown that the sperm–egg interaction is mediated by the vitelline-coat protein HrVC70 on the egg, and HrUrabin on sperm (Urayama et al., 2008). HrUrabin and its Ciona intestinalis orthologue CiUrabin are members of the GLIPR subfamily. HrUrabin and CiUrabin are highly expressed in testis (Urayama et al., 2008; Yamaguchi et al., 2011). Both contain a signal peptide, a N-terminal CAP domain and a GPI-anchor attachment site at the C-terminal domain (Urayama et al., 2008; Yamaguchi et al., 2011). Although the authors describe two CAP motifs, the HrUrabin and CiUrabin CAP domain actually contain all four CAP motifs (as described above). A classic example for involvement of the CAP domain in chemotaxis is the frog, Xenopus laevis, protein Allurin (Al-Anzi and Chandler, 1998; Olson et al., 2001). While originally described as a CRISP protein, it is actually a member of the GLIPR subgroup of CAPs. Allurin is a 21-kDa protein comprising two domains—a CAP and a hinge domain (Olson et al., 2001; Burnett et al., 2008). Allurin is expressed in the oviduct and localizes onto the outermost jelly layer. It is proposed that as eggs are shed into pond water, the gradient of allurin binds to sperm and guides them towards the site of the egg (Burnett et al., 2011; Burnett et al., 2012). Interestingly, allurin was also found to act as a mammalian sperm chemoattractant, thus raising the possibility that similar molecules, and functions, exist in mammals (Burnett et al., 2011; Burnett et al., 2012). Collectively, based on the invertebrate and vertebrate studies, it is proposed that the chemoattractive action of CAPs is mediated via the CAP domain. While poorly studied in mammals, CAP proteins are expressed in the female mammalian reproductive tract—CRISP1, CRISP2 and CRISP3 (Reddy et al., 2008; Ernesto et al., 2015; Evans et al., 2015). Of relevance here is that rat CRISP4 (called CRISP1 in the publication) has been proposed to act as a sperm chemoattractant (Ernesto et al., 2015). Moreover, the gradient of CRISP4 was found to stimulate sperm orientation and linear trajectories towards the egg (Ernesto et al., 2015). Although it has yet to be tested, the authors speculated that the chemotaxis action is mediated by the CRISP domain. CAP Superfamily Protein in Other Pathologies Although there is a notable expression bias to the male reproductive tract, CAP proteins are widely expressed across tissues in mammals (Reddy et al., 2008). In this section, we will discuss the potential roles of CAP superfamily proteins in non-fertility-related pathologies. CAPs in immune regulation As indicated throughout this review, there is emerging and almost overwhelming data to suggest that CAP proteins, and in particular the CAP domain, play a role in regulating the immune system. This has been most extensively explored in non-vertebrate CAPs including the venom allergen (Ag5), pathogenesis related-1 (Pr-1) and the bacterial SCP domain-containing proteins. These roles are summarized below. As described above, Pr-1 proteins have been shown to have anti-microbial functions (Gamir et al., 2017). Upon pathogenic invasion by Phytophthora infestans, plants upregulate Pr-1 protein production, which can bind and sequester the sterols within the pathogen, thus inhibiting its growth (Gamir et al., 2017). Moreover, recombinant Pr-1 protein from tomato (P14c) and tobacco (Pr-1a) inhibited the invasive pathogen Phytophthora brassicae at a concentration of 1.1 μM (Gamir et al., 2017). While this is an important finding, the same inhibitory effect could not be replicated with Aspergillus niger or Botrytis cinerea, which are sterol-prototroph fungal pathogens, thus raising some doubt over the anti-microbial function of Pr-1 proteins. As mentioned above, ASP proteins provide an additional example of the immune regulatory function of Pr-1 proteins, but they can also be used as a means for pathogens to subvert the host’s immunity to promote survival. ASP is also called activation-associated secreted protein and is secreted by canine hookworms (Anclystoma caninium) to avoid immune rejection. A. caninium produces CAP proteins (bacterial SCP-domain containing protein subfamily) including NIF, hookworm platelet inhibitor (HPI), ASP1 and ASP2 during the early stages of parasitic invasion (Moyle et al., 1994; Muchowski et al., 1994; Hawdon and Hotez, 1996; Hawdon et al., 1999). NIF is a 41-kDa glycoprotein which blocks the adhesion of neutrophils to vascular endothelial cells (Moyle et al., 1994). This is achieved by binding to CD11a and CD18 integrins (Muchowski et al., 1994; Lo et al., 1999), which play crucial roles in the mammalian innate immune system and inflammation (Moyle et al., 1994; Barczyk et al., 2010). Further, HPI—a CAP produced by A. caninum—was identified in a class of integrin receptor antagonists (Del Valle et al., 2003). Specifically, HPI can block the surface integrin receptors IIb/IIIa and GPIa/IIa resulting in the inhibition of platelet aggregation and adhesion (Del Valle et al., 2003). Similar platelet inhibitory activity was also reported in tablysin-15, a CAP protein in the saliva of the horsefly Tabanus yao (Xu et al., 2012). Tablysin-15 can also bind to integrins and to pro-inflammatory cysteinyl leukotrinenes with submicromolar affinities (Xu et al., 2012). Mouse CRISP3 protein has also been suggested to play a role in immune regulation (Pfisterer et al., 1996). CRISP3 is expressed in murine pre-B cell lines, bone marrow and spleen and its expression is regulated by the transcription factor OCT2 (Pfisterer et al., 1996). In mice, OCT2 is expressed during B cell development and is essential for the late phase of B cell differentiation (Corcoran and Karvelas, 1994; Hodson et al., 2016). As an example from mosquitoes, the expression levels of the cysteine-rich venom protein (venom allergen subfamily protein) CRVP379 is significantly upregulated during the infection of Aedes aegypti with dengue virus (DENV), a mosquito-borne flavivirus (Colpitts et al., 2011). Specifically, CRVP379 was highly upregulated on Day 1 and Day 14 of DENV infection (Bonizzoni et al., 2012). Of more direct relevance, in vivo and in vitro knockout down of CRVP379 resulted in significant reductions in viral infection and revealed a positive association between CRVP379 levels and DENV infections, indicating the requirement of CRVP379 to induce DENV infection in mosquito (Londono-Renteria et al., 2015). CRVP379 was shown to interact with prohibitin, a potential receptor for DENV in mosquitoes (Kuadkitkan et al., 2010; Londono-Renteria et al., 2015). CAPs in fibrosis Initially, CRISPLD2 (named late gestation lung 1, LGL1, at the time) was identified as a glucocorticoid-regulated mesenchymal secretory protein and shown to regulate lung branching and alveogenesis through mesenchymal–epithelial interactions (Himes et al., 2014). Later CRISPLD2 was implicated in inflammation. In foetal rat lungs, CRISPLD2 is secreted by fibroblasts at the tips of the budding alveoli (Nadeau et al., 2006). In agreement, CRISPLD2 was significantly downregulated in a rat bronchopulmonary dysplasia model (Nadeau et al., 2006). Bronchopulmonary dysplasia is a chronic lung disease in which infants require mechanical ventilation, oxygen, corticosteroid therapy and nutritional support (Wright and Kirpalani, 2011; Davidson and Berkelhamer, 2017). Recombinant CRISPLD2 was shown to directly affect migration of lung epithelial cells and had antagonistic effects on lipopolysaccharide-induced inflammation (Zhang et al., 2016). These data suggest that CRISPLD2 is a potential anti-inflammatory protein and that its suppression leads to an increased risk of lung inflammation (Wang et al., 2009; Zhang et al., 2016). The mechanism underpinning CRISPLD2 function is, however, still far from understood. GAPR was found to be elevated in the kidneys of mice with fibrosis, notably in the tubule epithelium, disrupted glomeruli and in the collecting ducts (Szyperski et al., 1998; Baxter et al., 2007). The underlying molecular mechanism of increased GAPR in fibrotic kidney is not known. However, in renal epithelial cell lines GAPR has been shown to induce trans-differentiation of epithelial cells to mesenchymal cells (Haverty et al., 1988). As such, it is believed that high levels of GAPR in kidneys can result in progression of fibrosis by increasing the activated fibroblasts via induction of the epithelial-to-mesenchymal transition (Baxter et al., 2007). GAPR was also found to regulate type I interferon signalling activation in response to toll-like receptor 4 (TLR4) in monocytes (Zhou et al., 2016). As noted throughout this review, numerous CAPs, including PI15, GAPR2 and CRISPLD2 subfamily proteins, have been implicated in the induction of epithelial-to-mesenchymal transitions (Baxter et al., 2007; Falak et al., 2014; Zhang et al., 2015). Collectively, these data indicate that this property is imparted by the CAP domain. CAPs in auto-immune diseases As a further reflection of the potential of CAP proteins to influence the immune system, several CAP proteins have been linked to autoimmune conditions, including in humans. For example, CRISP3 levels are significantly reduced in saliva of women with Sjögren’s syndrome (SS) compared to healthy controls (Laine et al., 2007). SS is a chronic autoimmune disorder characterized by the infiltration of lymphocytes into exocrine glands and B cell hyperactivation and is classically manifested in peri-menopausal women and associated with ocular and oral dryness (Skopouli et al., 2000; Mavragani and Moutsopoulos, 2014). Whether changes in CRISP3 expression are causal in SS or secondary to decreased circulating androgens that occur during the peri-menopausal period is yet to be determined. One, however, may feed into the other. Specifically, it is plausible that the natural decline in androgens that occurs during the peri-menopausal period (linked to decreasing oestrogen levels) leads to decreased CRISP3 promoter activation and decreased expression (Tapinos et al., 2002). This, in turn, may mean there is less CRISP3 available to induce a state of immune tolerance, similar to that described in section above. This hypothesis is yet to be tested directly. Additionally, it has been speculated that the CRISP3 gene can also be activated through an intermediary interaction between phorbol myristate acetate-activated peripheral blood lymphocytes and protein kinase C, in addition to androgens (Tapinos et al., 2002). As SS involves disruption of aquaporin-5 ion channel function, it has been proposed that the ICR of CRISP3 may regulate aquaporin-5 activity; however, this hypothesis has not been tested yet (Laine et al., 2007). It has also been suggested that CRISP3 may play a role in the aetiology of myasthenia gravis, a chronic autoimmune disorder of neuromuscular junctions (Berrih-Aknin et al., 2014). Specifically, a transcriptome analysis of peripheral blood mononuclear cells from patients with acetylcholine receptor-positive myasthenia gravis revealed that CRISP3 expression is significantly downregulated (Barzago et al., 2016). In contrast, CRISP3 expression is highly upregulated (~21-fold increase) in chronic pancreatitis (Friess et al., 2001), which is a recurrent autoinflammatory disease of the pancreas, which leads to severe exocrine and endocrine insufficiency (Lew et al., 2017). CRISP3 is localized to pancreatic acinar cells and is highly upregulated in chronic pancreatitis (Liao et al., 2003). The clinical relevance of this association is unknown. Further, CRISP3 is significantly upregulated (~10-fold increase) in peripheral blood samples from patients with multiple myeloma, a neoplastic plasma cell disease (Leich et al., 2013; Leng et al., 2018). CAPs in cancer CRISP3 and GLIPR1 have long been suggested to play a role in the development and progression of cancers. The expression of CRISP3 is highly upregulated in prostatic adenocarcinoma compared to healthy prostate tissue (Krätzschmar et al., 1996; Asmann et al., 2002; Ernst et al., 2002; Kosari et al., 2002). It has been speculated that CRISP3 plays a role in cancer progression by forming a complex with microseminoprotein-ß (MSMB) and that an imbalance in this binding promotes cancer progression (Udby et al., 2005; Ghasriani et al., 2009). MSMB is a protein with tumour inhibitory effects and proapoptotic activity in prostate cancer cell lines (Garde et al., 1999; Shukeir et al., 2003; Pathak et al., 2010). In malignant prostate, expression of MSMB is significantly downregulated and it is suggested that prostate cancer development may be affected by the amount of unbound CRISP3 (Chan et al., 1999). As such, CRISP3 has been suggested as a potential biomarker for prostate carcinomas (Kosari et al., 2002; Bjartell et al., 2007). The significance of CRISP3 in prostate malignancy is further supported by a 53-fold increase in CRISP3 mRNA expression associated with the TMPRSS2-ERG gene, a recurrent fusion gene found in the majority of prostate cancers (Iljin et al., 2006; Tomlins et al., 2008; Ribeiro et al., 2011). A direct role for CRISP3 in prostate cancer progression has not, however, been tested. In contrast to CRISP3, GLIPR1 is downregulated in human prostate cancer. Transfection of GLIPR1 into human and mouse prostate cell lines and viral delivery of Glipr1 cDNA in a prostate cancer model suppressed tumour growth and metastasis, and increased p53 regulated apoptosis (Ernst et al., 2002; Ren et al., 2002, 2006b; Satoh et al., 2003). Moreover, Glipr1-deficient mice were found to be more prone to the spontaneous initiation of prostatic tumour growth than wild-type animals (Li et al., 2008). The exact molecular function of GLIPR1 and its association with malignant prostate tumour remains uncharacterized. However, lines of evidence suggest that the over-expression of GLIPR1 promotes apoptosis of prostate cancer cells via activation of the JNK pathway, downregulation of c-Myc and suppression of aurora kinase A and targeting protein for Xenopus kinase-like protein 2 (Li et al., 2008, 2013). In addition, GLIPR1 has been proposed as a potential therapeutic marker in a range of other cancers (Sheng et al., 2016). Increased GLIPR1 expression is found in Wilms tumours, astrocytic, acute myeloid leukemia and melanomas (Rosenzweig et al., 2006; Chilukamarri et al., 2007; Xiao et al., 2011; Awasthi et al., 2013; Bier et al., 2013; Jacoby et al., 2014). GLIPR1 expression is significantly reduced in human lung cancer cells compared to normal cells and is believed to play a role in regulating lung tumorigenesis through tyrosine kinase receptors (Sheng et al., 2016). High levels of GLIPR1 were found in cisplatin-resistant human lung adenocarcinoma cells, which upon downregulation in vitro led to increased apoptosis. This downregulation also increases caspase-3, truncated poly (ADP-ribose) polymerase and significantly reduces B-cell lymphoma 2 protein expression in cisplatin-resistant cells (Gong et al., 2017). CAPs as proteases and protease inhibitors The function of CAPs as proteases was initially reported from the cone snail (Conus textile) CAP, Tex31 (Milne et al., 2003). Tex31 showed a proteolytic cleavage activity against the conotoxin TxVII pro-peptide and was suggested to be a novel substrate-specific protease (Milne et al., 2003). However, these findings were questioned when low level serine-protease-like activity was found in Mr30 (also referred to as GalCrisp)—a high sequence homology CAP protein from Conus marmoreus (Hansson et al., 2006; Qian et al., 2008). By contrast, others have proposed that CAP proteins have protease inhibitory functions. Frost and Engelhardt identified PI16 as a paracrine factor with potential cardiac function (Frost and Engelhardt, 2007). In animal disease models and humans, PI16 is significantly upregulated in diseased myocardium and inhibits cardiomyocytes growth in vivo and in vitro (Frost and Engelhardt, 2007). PI16 is secreted by cardiac fibroblasts and is proposed to inhibit the proteolytic cleavage of pro-chemerin and prevent pro-inflammatory activation of chemerin-sensitive cells via the CMKLR1 receptor (Regn et al., 2016). In support of this hypothesis, elevated levels of chemerin were observed in Pi16-deficient mice as compared to controls, and this was consistent with data reported in humans (Bozaoglu et al., 2010; Regn et al., 2016). Recombinant PI16 was also able to inhibit cathepsin K, a chemerin-activating protease, in a dose-dependent manner suggesting PI16 suppresses chemerin activation in the stressed myocardium (Regn et al., 2016). Splice variants of PI16 mRNA (PI16 isoforms 1–4) are also highly upregulated in human coronary artery endothelial cells following laminar shear stress and can inhibit endothelial migration via MMPs (Hazell et al., 2016). CAPs in reptile venoms Proteomic analysis has revealed that CRISPs constitute up to 3% of some snake venoms (copperhead snake, western diamondback rattlesnake, prairie rattlesnake and venomous pit viper) where they have been shown to induce paralysis of peripheral smooth muscle and induction of hypothermia in prey (Fry, 2005; Carregari et al., 2018). Based on the most widely accepted hypotheses of toxin evolution, the origin and evolution of toxicity in snake venom occurred via the recruitment of a non-venomous ‘body’ protein into the venom glands—in this case, modified salivary glands, which produce high concentrations of CRISPs (including in mammals). Subsequently, the gene was further modified through gene duplication and neofunctionalization events to increase toxicity (Lynch, 2007; Fry et al., 2009; Fry et al., 2012; Sunagar et al., 2012; Hargreaves et al., 2014). As detailed in Table I, reptilian CRISPs regulate a range of ion channels including calcium channels, potassium channels, ryanodine receptors and cyclic nucleotide-gated ion channels (Nobile et al., 1994, 1996; Yamazaki et al., 2002; Shikamoto et al., 2005; Wang et al., 2005, 2006, 2010; Peichoto et al., 2009; Adade et al., 2011). In addition to the ion channel regulatory roles, vCRISPs can regulate prey responses in other ways. For example, intraperitoneal injection of Bj-CRP from Bothrops jararaca (Pit viper) venom into mice induces an inflammatory response including the production of IL6 within the peritoneal cavity. It also induced generation of C3a, C4a and C5a anaphylatoxins of the human complement system (Lodovicho et al., 2016). The significance of snake CRISPs in immune regulation is further supported by the finding that natrin, from Naja atra (Chinese cobra), can regulate expression of vascular endothelial cell adhesion molecules including ICAM-1, VCAM-1 and E-selectin (Wang et al., 2010). ES-CRISP, a vCRISP from Echis carinatus sochureki (venomous viper), was identified as an anti-angiogenic protein which inhibits MAPK ERK1/2 signalling through direct interactions with endothelial cells, followed by internalized in a granule-like manner (Lecht et al., 2015). Mechanistically, once internalized into the cytoplasm, ES-CRISP downregulates transforming growth factor-ß expression, which is a positive angiogenesis regulator across various cancers (Pozzi et al., 2000; Bikfalvi, 2004; Saunier and Akhurst, 2006; Lecht et al., 2015). CAPs in treating disease Just as the induction of CAP proteins may induce disease, their regulation may offer opportunities for therapy. One such example is crovirin, a snake venom CRISP from Crotalus viridis (rattle snake) (Adade et al., 2014). Crovirin has anti-parasitic activity against the developmental stages of Trypanosoma cruzi (Adade et al., 2011), the causal pathogen in Chagas disease, in its insect vectors. Crovirin has low toxicity to host cells and mice, and non-lethal doses were shown to significantly inhibit Trypanosoma cruzi and Leishmania amazonensis proliferation (Utkin and Osipov, 2007; Peichoto et al., 2009; Adade et al., 2014). Together, these studies show crovirin to have anti-parasitic, trypanocidal and leishmanicidal activity (Adade et al., 2011, 2014). Conclusion Within this review, we have attempted to unify data from the full spectrum of CAP superfamily proteins, with the intent of highlighting commonality of function across members spread throughout the biological kingdoms. We highlight, and elaborate on, the stepwise evolutionary process proposed by Abraham and Chandler (2017), as to how the single domain proteins in prokaryotes evolved into multi-domain proteins in complex eukaryotes. While large gaps in the field’s knowledge of CAP biology remain, data generated in the last decade in particular has solidified their place as critical players in biological processes across all kingdoms. Data indicate that CAP domains are critical regulators of the immune system, sterol/lipid binding domains, are likely protease inhibitors and mediators of epithelial-to-mesenchymal transitions. Similarly, the C-terminal extensions have earned their place as key regulators of ion channel function, in both venomous reptiles and mammalian male fertility. Much, however, needs to be learned about the function of the C-terminal extension in other CAPs subfamilies. Mutations in human CAP genes have also been clearly linked to the aetiology of diseases, including male infertility, neurological disorders, audio-vestibular dysfunction, heart disease and bronchopulmonary dysplasia, and their dysregulation linked to other conditions including prostate and lung cancer, chronic pancreatitis, glomerulonephritis and a range of autoimmune conditions. The strong expression bias of CAP proteins to the male reproductive tract in both mammals and insects, and their proposed functions across multiple aspects of fertility, remains an enigma. While all functional and evolutionary data point to a role in enhancing gamete function and survival in the female reproductive tract, many of these functions are yet to be directly tested. These data are also confounded, in part, by the imprecise use of nomenclature. We hope this review will serve as a framework for future studies to define the full spectrum of CAP biology. Acknowledgements The authors acknowledge Anne O’Connor for her review of this manuscript. Authors’ roles A.S.G. and M.O’.B. conducted the literature search and wrote the initial draft of the manuscript. A.S.G. and J.H. prepared the figures. A.S.G. and D.G.C. carried out the phylogenetic analysis. All authors contributed to reviewing and editing of the final manuscript. Funding Australian Research Council grant (DP200100659 to M.K.O.B.) Conflict of interest The authors declare no conflict of interest. References Abraham A , Chandler DE. Tracing the evolutionary history of the CAP superfamily of proteins using amino acid sequence homology and conservation of splice sites . J Mol Evol 2017 ; 85 : 137 – 157 . Google Scholar Crossref Search ADS PubMed WorldCat Acevedo JJ , Mendoza-Lujambio I, La Vega-Beltrán JL De, Treviño CL, Felix R, Darszon A. KATP channels in mouse spermatogenic cells and sperm, and their role in capacitation . Dev Biol 2006 ; 289 : 395 – 405 . 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TI - The functions of CAP superfamily proteins in mammalian fertility and disease
JF - Human Reproduction Update
DO - 10.1093/humupd/dmaa016
DA - 2020-09-01
UR - https://www.deepdyve.com/lp/oxford-university-press/the-functions-of-cap-superfamily-proteins-in-mammalian-fertility-and-kRfPJshk8u
SP - 689
EP - 723
VL - 26
IS - 5
DP - DeepDyve
ER -