Antimicrobial peptides (Homo sapiens)

LL-37 has direct microbicidal activities against Gram-positive bacteria (S. aureus, Group A Streptococcus, Bacillus megaterium), Gram-negative bacteria (E. coli, P. aeruginosa, Salmonella minnesota) and fungi such as C. albicans (Yang D et al. 2000; Nagaoka I et al. 2005; Braff MH et al. 2005; Wang G et al. 2012). LL-37 also has antiviral activities against herpes simplex virus, HIV-1, and vaccinia virus (Yasin B et al. 2000; Steinstraesser L et al. 2005; Howell MD et al. 2006; Gordon YJ et al. 2005). LL-37 may have the potential to prevent sepsis or septic shock associated with pathogenic bacterial infection by inhibiting the release of toxic components such as LPS and lipoteichoic acid (LTA) that cause excess tissue damage and inflammation (Larrick JW et al. 1995)

LL-37 expression was found to correlate with an activation of TLR2, TLR4 and TLR9 signaling pathways in M. tuberculosis-stimulatied human monocyte-derived macrophages, alveolar macrophages and neutrophils (Rivas-Santiago et al. 2008).

PGRPs are thought to kill bacteria by interacting with cell wall peptidoglycan and by inducing lethal stress response in bacteria, rather than a hydrolysis of peptidoglycan or permeabilizing bacterial membranes as other antibacterial peptides do (Lu X et al. 2006; Wang M et al. 2007; Cho S et al. 2007; Kashyap DR et al. 2011, 2014). In Gram-positive bacteria, including B. subtilis, PGLYRP1, 3 & 4 were found to enter cell wall at the site of daughter cell separation during cell division and to bind to cell wall peptidoglycan in the vicinity of cell membrane (Kashyap DR et al. 2011). However, this binding did not inhibit the extracellular transglycosylation or transpeptidation steps in peptidoglycan synthesis (Kashyap DR et al. 2011), which are well-known targets for bactericidal antibiotics. Instead, this interaction of PGRP proteins and peptidoglycan activated the B. subtilis envelope stress response CssR‑CssS two-component system that detects and disposes of misfolded proteins exported out of bacterial cells. This activation resulted in bacterial membrane depolarization, cessation of intracellular peptidoglycan, protein, RNA, and DNA synthesis, and production of toxic hydroxyl radicals, which were responsible for bacterial death (Kashyap DR et al. 2011). PGRPs were shown to kill Gram-negative bacteria (E. coli) by binding to their outer membrane and activating the functionally homologous CpxA-CpxR two‑component system (Kashyap DR et al. 2011). Furthermore, genome expression arrays, qRT-PCR, and biochemical tests showed that PGRPs kill both E. coli and B. subtilis by inducing oxidative, thiol, and metal stress (Kashyap DR et al. 2014).

There is also emerging evidence that PGLYRP1 can function as a receptor agonist or antagonist for human cells. Human PGLYRP1 (multimerized or complexed with peptidoglycan) binds to and stimulates triggering receptor expressed on myeloid cells-1 (TREM-1), a receptor present on neutrophils, monocytes and macrophages that induces production of pro-inflammatory cytokines (Read CB et al. 2015). Moreover, PGLYRP1 and its complex with 70-kDa heat shock protein (Hsp70) bind to the tumor necrosis factor receptor-1 (TNFR1, which is a death receptor). The PGLYRP1-Hsp70 complex induces a cytotoxic effect via apoptosis and necroptosis (Yashin DV et al. 2015, 2016), which accounts for the tumor cytotoxicity of PGLYRP1-Hsp70 complexes secreted by cytotoxic lymphocytes (Sashchenko LP et al. 2004). By contrast, free PGLYRP1 acts as a TNFR1 antagonist, by binding to TNFR1 and inhibiting its activation by PGLYRP1-Hsp70 complexes.

Both S100A8 and S100A9 belong to the S100 family of helix-turn-helix (EF-hand) calcium Ca(2+)-binding proteins. S100 proteins are involved in a wide range of cellular functions (Donato R et al. 2013; Zackular JP et al. 2015; Vogl et al. 2007). Within cells, S100 proteins are involved in aspects of regulation of proliferation, differentiation, apoptosis, Ca(2+) homeostasis, inflammation and migration/invasion (Donato R et al. 2013). During infection, certain S100 proteins can be secreted or released by cells to act as damage-associated molecular patterns (DAMPs) and interact with pattern recognition receptors to modulate inflammatory responses (Foell D et al. 2007; Vogl et al. 2007). In addition, these inflammatory S100 proteins have antimicrobial function by sequestering essential transition metals from bacteria, preventing their growth (Zackular JP et al. 2015). The fundamental structural unit of S100 proteins is a highly integrated antiparallel dimer (Potts BC et al. 1995; Heizmann CW et al. 2002; Brodersen DE et al. 1999; Moroz OV et al. 2009; Gagnon DM et al. 2015). All S100 proteins form this structure as homodimers. S100A8 and S100A9 are unique among all members of the S100 family because they preferentially form a heterodimer. Calprotectin (S100A8:S100A9) and other S100 proteins are Ca(2+)-activated regulators (Brophy MB et al. 2012; Donato R et al. 2013). Inside the cell, where the basal level of Ca(2+) is in the nanomolar range, S100 proteins can serve as a sensor of Ca(2+)-mediated signals. In the extracellular milieu, S100 proteins are perpetually (Ca2+)-bound because Ca(2+) concentration is in the millimolar range. Ca(2+) is also known to stimulate formation of higher order oligomers of S100 proteins, including S100A8/S100A9 tetramers (Leukert N et al. 2006; Korndörfer IP et al. 2007). Upon dimerization S100A8 and S100A9 form two metal binding sites at the dimer interface, both of which can bind to Zn(2+) with high affinity (Kd Zn(2+) about 10e-9 M) (Damo SM et al. 2013; Brophy MB et al. 2013). A chelation of Mn(2+) involves a single binding site (Kd Mn(2+) around 10e-7 - 10e-8 M) (Damo SM et al. 2013; Hayden JA et al. 2013; Gagnon DM et al. 2015).

Thus, calprotectin S100A8:S100A9 inhibits bacterial growth by targeting transition metals and sequestering these metals in a process referred to as nutritional immunity.

Structural studies revealed that S100A7 functions as (Ca2+)-bound homodimer, which can load two Zn2+ ions at symmetrically disposed sites across the dimer interface using residues His-86 and His-90 from one subunit and residues His-17 and Asp-24 from the other (Brodersen DE et al. 1999). Binding of Zn2+ is believed to stabilize the dimer and potentially mediate S100A7 function during infection (Brodersen DE et al. 1999).

S100A7 and its paralog, S100A7A (S100A15 or koebnerisin) display 93% sequence identity (Wolf R et al. 2011; Murray JI et al. 2012). Human S100A7A (S100A15) showed antimicrobial activity against E. coli (Büchau AS et al. 2007). Moreover, structural and solution binding studies revealed similar affinities of zinc ion for S100A7 and S100A7A (S100A15) (Murray JI et al. 2012). Though the Reactome project describes psoriacin (S100A7) and koebnerisin (S100A7A) as antimicrobial proteins with the metal-chelating properties, additional studies are needed to more fully define the contribution of S100A7 and S100A7A to nutritional immunity.

Particularly high expression of S100A7 and S100A7A was observed in inflamed psoriatic lesions, which are characterized by disturbed epidermal differentiation and inflammation (Madsen P et al. 1991). Circulating leukocytes (PBMCs) of patients with psoriasis produced increased levels of koebnerisin and psoriasin compared to healthy individuals (Batycka-Baran A et al. 2015). Both S100A proteins further acted as 'alarmins' on PBMC to induce proinflammatory cytokines implicated in the pathogenesis of psoriasis, such as IL-1beta, TNFalpha, IL6 and IL8 (Batycka-Baran A et al. 2015). However, inflammatory activities of S100A7 and S100A7A were found to serve distinct roles in epithelial homeostasis, inflammation, and cancer (Hattinger E et al. 2013; Wolf R et al. 2011; Murray JI et al. 2012). S100A7 signals through the receptor for advanced glycation products (RAGE) in a zinc-dependent manner, while S100A15 signals through a yet unidentified G-protein coupled receptor in a zinc-independent manner (Wolf R et al. 2011; Murray JI et al. 2012). Apart from inflammatory skin diseases an elevated exression of S100A7 was found in several epithelial cancers such as squamous cell carcinoma (SCC) of the skin, bladder, lung as well as in in situ ductal breast carcinoma (Celis JE et al. 1996; Al-Haddad S et al. 1999; Emberley ED et al. 2004; Moubayed N et al. 2007; Liu G et al. 2015; Qi Z et al. 2015).

Both S100A8 and S100A9 belong to the S100 family of helix-turn-helix (EF-hand) calcium Ca(2+)-binding proteins. S100 proteins are involved in a wide range of cellular functions (Donato R et al. 2013; Zackular JP et al. 2015; Vogl et al. 2007). Within cells, S100 proteins are involved in aspects of regulation of proliferation, differentiation, apoptosis, Ca(2+) homeostasis, inflammation and migration/invasion (Donato R et al. 2013). During infection, certain S100 proteins can be secreted or released by cells to act as damage-associated molecular patterns (DAMPs) and interact with pattern recognition receptors to modulate inflammatory responses (Foell D et al. 2007; Vogl et al. 2007). In addition, these inflammatory S100 proteins have antimicrobial function by sequestering essential transition metals from bacteria, preventing their growth (Zackular JP et al. 2015). The fundamental structural unit of S100 proteins is a highly integrated antiparallel dimer (Potts BC et al. 1995; Heizmann CW et al. 2002; Brodersen DE et al. 1999; Moroz OV et al. 2009; Gagnon DM et al. 2015). All S100 proteins form this structure as homodimers. S100A8 and S100A9 are unique among all members of the S100 family because they preferentially form a heterodimer. Calprotectin (S100A8:S100A9) and other S100 proteins are Ca(2+)-activated regulators (Brophy MB et al. 2012; Donato R et al. 2013). Inside the cell, where the basal level of Ca(2+) is in the nanomolar range, S100 proteins can serve as a sensor of Ca(2+)-mediated signals. In the extracellular milieu, S100 proteins are perpetually (Ca2+)-bound because Ca(2+) concentration is in the millimolar range. Ca(2+) is also known to stimulate formation of higher order oligomers of S100 proteins, including S100A8/S100A9 tetramers (Leukert N et al. 2006; Korndörfer IP et al. 2007). Upon dimerization S100A8 and S100A9 form two metal binding sites at the dimer interface, both of which can bind to Zn(2+) with high affinity (Kd Zn(2+) about 10e-9 M) (Damo SM et al. 2013; Brophy MB et al. 2013). A chelation of Mn(2+) involves a single binding site (Kd Mn(2+) around 10e-7 - 10e-8 M) (Damo SM et al. 2013; Hayden JA et al. 2013; Gagnon DM et al. 2015).

Thus, calprotectin S100A8:S100A9 inhibits bacterial growth by targeting transition metals and sequestering these metals in a process referred to as nutritional immunity.

PGRPs are thought to kill bacteria by interacting with cell wall peptidoglycan and by inducing lethal stress response in bacteria, as opposed to the hydrolysis of peptidoglycan or permeabilizing bacterial membranes seen with other antibacterial peptides (Lu X et al. 2006; Wang M et al. 2007; Cho S et al. 2007; Kashyap DR et al. 2011, 2014). In Gram‑positive bacteria, including B. subtilis, PGLYRP1, 3 & 4 were found to enter the bacterial cell wall at the site of daughter cell separation during cell division and to bind to cell wall peptidoglycan in the vicinity of the cell membrane (Kashyap DR et al. 2011). Binding of PGLYRP3 to peptidoglycan induces a structural change in PGLYRP3 that locks peptidoglycan in the protein's bindings groove (Guan R et al. 2006). However, this binding did not inhibit the extracellular transglycosylation or transpeptidation steps in peptidoglycan synthesis (Kashyap DR et al. 2011), which are well-known targets for bactericidal antibiotics. Instead, this interaction of PGRP proteins with peptidoglycan activated the B. subtilis envelope stress response two‑component system CssR‑CssS that detects and disposes of misfolded proteins exported out of bacterial cells. This activation resulted in bacterial membrane depolarization, cessation of intracellular peptidoglycan, protein, RNA, and DNA synthesis and production of toxic hydroxyl radicals, which were responsible for bacterial death (Kashyap DR et al. 2011). In Gram‑negative bacteria (E. coli), PGRPs were found to bind the bacterial outer membrane activating the functionally homologous CpxA‑CpxR two‑component system (Kashyap DR et al. 2011). Genome expression arrays, qRT‑PCR, and biochemical tests showed that PGLYRP3 & 4 kill both E. coli and B. subtilis by inducing oxidative, thiol, and metal stress (Kashyap DR et al. 2014).

DCD(63-109) shows a broad spectrum of antimicrobial activity against Gram-positive (Staphylococcus aureus, Enterococcus faecalis, Staphylococcus epidermidis, Listeria monocytogenes) and Gram-negative bacteria (Escherichia coli, Pseudomonas putida, Salmonella typhimurium) as well as Candida albicans (Cipakova I et al. 2006, Lai YP et al. 2005, Schittek B et al. 2001, Steffen H et al. 2006, Vuong C et al. 2004). The activity of the DCD(63-109) was maintained over a broad pH range and in high salt concentrations that resembled the conditions in human sweat (Schittek B et al. 2001).

RNase A family is a vertebrate-specific gene family (Goo SM & Cho S 2013). Members of RNase A family share specific elements of sequence homology, a unique disulfide-bonded tertiary structure, and the ability to hydrolyze polymeric RNA (Beintema JJ & Kleineidam RG 1998; Rosenberg HF 2008). Eight catalytically active members are found in humans: RNase1 (pancreatic RNase), RNase2 (eosinophil derived neurotoxin/EDN), RNase3 (eosinophil cationic protein/ECP), RNase4, RNase5 (angiogenin), RNase6, RNase7 (skin-derived RNase), and RNase8 (divergent paralog of RNase7) (Sorrentino S 2010). Analysis of human genome sequence has revealed the existence of five additional RNases named as RNases 9-13, although they appear to lose enzymatic activity (Devor EJ et al. 2004; Castella S et al. 2004; Cho S et al. 2005). All human RNase A family members encode relatively small polypeptides of 14 to 16kDa containing signal peptides of 20 to 28 amino acids for protein secretion. Mature RNases contain 6 to 8 cysteine residues that are crucial to hold the overall tertiary structure (Sorrentino S 2010). Apart from the ribonuclease activity the RNase A family members have been implicated in a wide variety of biological actions including antipathogen and immunomodulatory activities (Harder J & Schroder JM 2002; Rudolph B et al. 2006; Boix E et al. 2008; Boix and Nogués, 2007; Spencer JD et al. 2011; Becknell B et al. 2015; Rosenberg HF 2015). Evidence of antimicrobial properties displayed by distantly related members ascribed to the family an ancestral role in host defence (Pizzo E & D’Alessio G 2007; Rosenberg HF et al. 2008).

RNase3, RNase6 and RNase7 have been identified as the most potent human antibacterial ribonucleases with a broad antimicrobial action against Gram-positive and Gram-negative bacteria (Pulido D et al. 2013, 2016; Zhang J et al. 2003; Boix E et al. 2008; Torrent M et al. 2010). Mutagenesis analysis revealed that ribonuclease-inactive RNase7 protein exhibited similar anti-microbial activity against P. aeruginosa, E. faecium and E. coli as the wild-type protein suggesting that RNase7 may kill bacteria independently of its ribonuclease catalytic activity (Huang YC et al. 2007; Koten B et al. 2009). Similar results were reported on microbicidal effect of ribonuclease-inactive RNase3 and 6 proteins against S. aureus (Rosenberg HF 1995; Pulido D et al. 2016a). Being cationic proteins with a high pI, RNase3, 6 and 7 interact with anionic components of biological membranes (Zhang J et al. 2003; Boix E et al. 2008; Torrent M et al. 2010; Boix E et al. 2012; Pulido D et al. 2016a). RNase3, 6 and 7 present, respectively, a high number of either Arg, His or Lys surface-exposed residues that may contribute to their distinct bactericidal mechanisms of action (Torrent M et al. 2010; Prats-Ejarque G et al. 2016). RNase3 displays a membrane disruption capacity that is dependent on both surface exposed hydrophobic and cationic residues. RNase3 can bind and partially insert into the lipid bilayers, promoting its aggregation and final lysis, following a carpet-like mechanism. The RNase3 agglutination process precedes the bacterial death and lysis event. The antimicrobial properties of the RNase6 are comparable to its RNase3 homolog and correlate to the bacterial cell damage and agglutination activities (Pulido D et al. 2016a). In contrast, RNase7 has no significant membrane aggregation capacity (Torrent M et al. 2010). RNase7 binds and permeabilizes the bacterial membrane displaying a much higher leakage capacity compared to RNase3 (Torrent M et al. 2010; Huang YC et al. 2007). Membrane permeabilization by RNase7 required four clustered lysine residues but no catalytic residues (Huang YC et al. 2007). Binding to PGN and LPS has been reported for RNases 3 and 7 (Torrent M et al. 2010; Pulido D et al. 2016b). Studies using a battery of progressively truncated LPS-defective E. coli strains correlated the LPS interaction with the protein cell agglutination and bactericidal activities (Pulido D et al. 2012). Further work indicated that RNase3 and RNase 6 high cell agglutination activity towards Gram negative species is retained by their respective N-terminus peptides (Torrent M et al. 2012, 2013; Pulido D et al. 2016c). In particular, the RNase3 N-terminus encompasses a specific patch (Y33-R36) required for LPS binding and an hydrophobic aggregation prone region (A8-I16) that mediates the protein self amyloid- like aggregation and promotes the cell death.

Mutations in the gene encoding ATP7A results in a severe copper-deficiency known as Menkes disease (Kaler SG 2011).

Secreted INTL1 functions as a disulfide-linked trimer (Tsuji S et al., 2001; Tsuji S et al., 2007; Wesener DA et al., 2015).

GNLY is synthesized as a 15-kDa molecule and then proteolytically cleaved at the amino and carboxyl termini to produce a 9-kDa form (Pena SV et al. 1997; Hanson DA et al. 1999). The 9 -kDa form of GNLY is confined to cytolytic granules that are directionally released by receptor-mediated granule exocytosis following target cell recognition (Hanson DA et al. 1999; Clayberger C et al 2012). In contrast, the 15-kDa form is constitutively secreted from distinct granules that lack perforin and granzyme (Clayberger C et al 2012). The 9-kDa GNLY exhibits cytolytic activity on the numerous microbes ranging from extracellular and intracellular bacteria to fungi and parasite (Stenger S et al. 1998; Ernst WA et al. 2000). GNLY kills Mycobacterium tuberculosis, the causative agent in tuberculosis and Plasmodium falciparum, a cause of malaria (Stenger S et al. 1998; Farouk SE et al. 2004). Alongside its ability to kill bacteria, fungi, and parasites, GNLY can block viral replication and trigger apoptosis in infected cells (Hata A et al. 2001).

Besides its direct antimicrobial activity, GNLY shows tumoricidal activity by inducing apoptosis in tumor cells. Both 9-kDA and 15-kDA forms of GNLY may also function as chemoattractants for T lymphocytes, monocytes and other inflammatory cells and activates the expression of a number of cytokines (Deng A et al. 2005; Castiello L et al. 2011).

HTNs exhibited antibacterial activity in vitro against various bacteria, including S. mutans, P. gingivalis, A. actinomycetemcomitans, P. aeruginosa and St. aureus (MacKay BJ et al. 1984; Murakami Y et al. 1991; Payne JB et al. 1991; Nishikata MH et al. 1991; Sajjan US et al. 2001; Murakami Y et al. 2002; Giacometti A et al. 2005; Welling MM et al. 2007). HTNs were also active against complex mixtures of bacteria, such as those present in saliva and plaque (Helmerhorst EJ et al. 1999). The antibacterial activity of HTNs is thought to rely on electrostatic interactions of cationic HTNs with anionic phospholipids, such as phosphatidylglycerol and cardiolipin on the bacterial cell surface (De Smet K & Contreras R 2005). HTNs have also been shown to possess a fungicidal activity (Oppenheim FG et al. 1988; Troxler RF 1990). HTN5 (a product of HTN3 gene) is the most potent among all histatin family members with regard to its antifungal activity against C.albicans and C.neoformans (Xu T et al. 1991; Tsai H & Bobek LA 1997; Helmerhorst EJ et al. 2001).

It is believed that the electrostatic interaction between the cationic GNLY and the negatively charged microbial cell surface increases the ability of GNLY to fold into amphipathic conformation which can disrupt microbial membranes resulting in cytolysis by osmotic shock (Wang Z et al. 2000; Ernst WA et al. 2000; Anderson DH et al. 2003; Barman H et al. 2006).

The functional role of HTNs peptides in vivo involves prevention of the oral overgrowth of Candida albicans in oropharyngeal candidiasis (Jainkittivong A et al. 1998; Khan SA et al. 2013). HTNs display candidacidal and candidastatic activities in vitro against Candida albicans, Candida glabrata, Candida dubliniensis, Candida krusei, Saccharomyces cerevisiae, Cryptococcus neoformans and Neurospora crassa (Tsai H & Bobek LA 1997, 1998; Oppenheim FG et al. 1988; Xu T et al. 1991; Rayhan R et al. 1992; Helmerhorst EJ et al. 1999; van't Hof W et al. 2000; Fitzgerald DH et al. 2003). Histatin 5 shows the highest anticandicidal activity of the family in vitro at physiological concentrations found in saliva (15–50 microM) (Xu T et al. 1991). The candidacidal activity of histatin 5 results from a multistep molecular mechanism involving the recognition and binding of the peptide to the yeast cell wall proteins Ssa1/2p followed by the peptide internalization (Edgerton M et al. 1998; Li XS et al. 2003; Sun JN et al. 2008). Histatin 5 internalization is required for fungicidal activity in Candida species (Li XS et al. 2006; Jang WS et al. 2010). Ultimately, histatin 5 is transported into the cell through the fungal polyamine transporters Dur3 and Dur31 in an energy-dependent process (Kumar R et al. 2011; Tati S et al. 2013). Histatin 5- mediated killing involves interaction with the fungal mitochondrial membrane (Helmerhorst EJ et al. 1999; Gyurko C et al. 2000; 2001). Interference with mitochondrial respiratory machinery can lead to generation of reactive oxygen species (ROS) and ATP release (Helmerhorst EJ et al. 2001; Gyurko C et al. 2000; 2001). The killing of C. albicans is accompanied by the release of intracellular potassium ions and the Trk1 potassium channel is critical (Pollock JJ et al. 1984; Baev D et al 2004; Vylkova S et al. 2006). Histatine 5 has also been shown to bind various metals in saliva namely, Zn, Cu, Fe and Ni that can modulate the peptide candidacidal properties (Melino S et al. 1999; 2014; Puri S et al. 2015).

Antineutrophil cytoplasmic autoantibodies against BPI (BPI-ANCA) have been found in diseases of different etiologies, such as cystic fibrosis, TAP deficiency or inflammatory bowel diseases (Walmsley RS et al. 1997; Schultz H et al. 2004; Schinke S et al. 2004; Aichele D et al. 2006). The presence of BPI-ANCA has been shown to correlate with the chronic or profuse exposure of the host to Gram-negative bacteria and their endotoxin (Aebi C et al. 2000; Carlsson M et al. 2003; Schultz H et al. 2003; Schultz H 2007).

Several CHGA-derived peptides such as vasostatin-1 (CHGA(19-94) ) and catestatin (CHGA(370-390)) display antimicrobial activities against bacteria, fungi and yeasts (Lugardon K et al. 2000; Briolat J et al. 2005; Radek KA et al. 2008; Aslam R et al. 2013; Shooshtarizadeh P et al. 2010). These peptides are found in biological fluids involved in defence mechanisms (human serum and saliva) and in supernatants of stimulated human neutrophils (Lugardon K et al. 2000; Briolat J et al. 2005). In addition, catestatin (CHGA(370-390) exhibits antimicrobial activity against skin pathogens suggesting a function in cutaneous antimicrobial defense (Radek KA et al. 2008). Biophysical and structural analysis of human catestatin and bovine cateslytin suggests that cationic CHGA-derived peptides interact with anionic phospholipids on the bacterial surface (Sugawara M et al. 2010; Jean-Francois F et al. 2008). However, It remains to be clarified whether catestatin functions as a pore-forming or cell-penetrating agent.

BPIFs are subdivided into two groups, the long (BPIFB or LPLUNC) and short (BPIFA or SPLUNC) proteins. The BPIFB (LPLUNC) proteins have sequence and structure homology to both the LPS-binding N-terminus and the C-terminus of BPI, which is responsible for its opsonization activity, and BPIFA (SPLUNC) have homology to only the N-terminal half of BPI (....). It is proposed that the BPIF proteins may function in an antimicrobial manner, however, showed a high degree of conservation of genomic organization and of exon sizes. all show the same conservation of two cysteine residues which form a critical disulphide bond (Beamer et al. 1997) The function of the BPIF family of proteins has yet to be elucidated, but due to their predicted structure and their similarity in gene location to known LPS binding proteins, BPI and LBP, it is proposed that they function in an antimicrobial manner may function in the innate immune system, against gram negative bacterial LPS, either by acting directly against bacteria in a bactericidal manner, initiating an immune response or as an anti-toxin, by reducing the inflammatory response.All members of this family appear to have the ability to bind to and transfer lipid molecules (Bingle and Craven, 2003). BPIFA1 (SPLUNC1) is secreted in the epithelium of the upper airways, where it coats the surface of the epithelium and cilia, but significantly greater expression is seen in the submucosal cells and ducts of glands associated with the upper airways (Campos MA et al. 2004; Di YP et al. 2003; Bingle L et al. 2005; Bingle L & Bingle CD 2011). BPIFA1 is considered to contribute to mucosa immunity protecting the upper airway from infections. The antibacterial properties BPIFA1 have been demonstrated with regard to Staphylococcus aureus, Streptococcus, Pseudomonas, Mycoplasma pneumonia and Klebsiella pneumonia (Di YP 2011; Gally F et al. 2011; Tsou YA et al. 201; Liu Y et al. 2013). It is also considered to have antibiofilm functionality through its ability to modulate surface tension of airway fluids (Gakhar L et al. 2010; Bartlett JA et al. 2011; Liu Y et al. 2013). In addition, BPIFA1 negatively regulates epithelial sodium channel (ENaC) function, affecting the height of the airway surface liquid, which is essential for adequate mucociliary clearance (Garcia-Caballero A et al. 2009; Hobbs CA et al. 2013; Tarran R & Redinbo MR 2014). Furthermore, BPIFA1 also shows immunomodulatory properties in different mouse models of acute airway inflammations (). The high level of BPIFA1 present in the airways under basal conditions fluctuating rapidly in response to environmental stress and upon airway inflammation. The fluctuations in the BPIFA1 levels are thought to help the local milieu in optimizing protective functions in the airways (Bingle L et al. 2007; Britto CJ and Cohn L 2015). SPLUNC2 is expressed in serous cells of the major salivary glands and in minor mucosal glands mainly secreted from the upper airway.

EPPIN has two potential protease inhibitory domains: a whey acid protein (WAP)-type four disulfide core (WFDC) domain and a Kunitz domain (Richardson RT et al. 2001; McCrudden MT et al. 2008). While both domains of EPPIN are involved in the protein's antibacterial activity, only the Kunitz domain is required for selective protease inhibition (McCrudden MT et al. 2008).

PI3 is constitutively expressed in a number of epithelial barriers that are constantly exposed to foreign antigens and pathogens, including skin, airway, and intestinal mucosa (Pfundt R et al. 1996). PI3 can be transcriptionally upregulated at various sites of inflammation by LPS or pro-inflammatory cytokines such as IL1B and TNF (Sallenave JM et al. 1994; Alkemade JA et al. 1994; Pfundt R et al. 2000, Simpson AJ et al. 2001).

Structural analysis of LEAP2 revealed a compact central core stabilized by two disulfide bonds between Cys17-28 and Cys23-33 and a network of hydrogen bonds (Henriques ST et al. 2010; Hocquellet A et al. 2010). The central core of LEAP2 contains the majority of the arginine and lysine residues, which are clustered to form an extended, positively-charged patch on the surface of the molecule (Henriques ST et al. 2010). The terminal segments of LEAP2 are relatively unstructured and contain the majority of the hydrophobic residues (Henriques ST et al. 2010). Membrane-affinity studies show that LEAP2 membrane binding is governed by electrostatic attractions, which are sensitive to ionic strength. Truncation studies found that the C-terminal region of LEAP2 is irrelevant for both membrane binding and antimicrobial activity, whereas the N-terminal (hydrophobic domain) and core regions (cationic domain) are essential (Henriques ST et al. 2010).

LEAP2 showed antimicrobial activity against Bacillus subtilis at low ionic strength (Henriques ST et al. 2010). The inability of LEAP2 to inhibit B. subtilis growth at physiologically relevant salt concentration is consistent with a proposed electrostatic contribution to membrane binding (Henriques ST et al. 2010). Other bacteria such as Gram-positive Bacillus megaterium, Staphylococcus carnosus, Micrococcus luteus and Gram-negative Neisseria cinerea are also sensitive to treatment with LEAP2 (Krause A et al. 2003). Furthermore, the native and reduced forms of LEAP2 show similar membrane affinity and antimicrobial activities; this suggests that disulfide bonds are not essential for bactericidal activity (Henriques ST et al. 2010; Hocquellet A et al. 2010). LEAP2 did not affect the growth of Escherichia coli and S. aureus at physiological or low ionic strength (Henriques ST et al. 2010). LEAP2 was found to be inactive against Pseudomonas fluorescens (Krause A et al. 2003). The antimicrobial potential of LEAP2 against E. coli, B. subtilis, B. megaterium, and M. luteus was significantly lower than that of cathelicidin LL37 (Hocquellet A et al. 2010). Further study is needed to clarify the antimicrobial activity of LEAP2.

LEAP2 is highly conserved throughout vertebrates, particularly in mammals suggesting that LEAP2 has additional physiological functions that remain to be elucidated (Krause A et al. 2003; Zhang YA et al. 2004; Townes CL et al. 2009; Henriques ST et al. 2010).

• attacking membrane-associated (E. coli) or capsule proteins (S.pneumonia), which leads to loss of membrane integrity (Belaaouaj A et al. 2000; Standish AJ & Weiser JN 2009)
• processing host immune proteins to generate antimicrobial peptides that can directly kill bacteria (Sorensen OE et al. 2001)
• targeting and inactivating bacterial virulence factors to attenuate bacteria (Weinrauch Y et al. 2002; Lopez-Boado YS et al. 2004)

Many pathogens such as Streptococcus pneumoniae have evolved lysozyme resistance to prevent peptidoglycan hydrolysis. The primary mechanism for lysozyme resistance in both Gram-positive and Gram-negative organisms appears to be direct modification of peptidoglycan; however, modification of other cell wall-linked components, such as teichoic acid, may also contribute to resistance (Amano K & Williams JC 1983; Bera A et al. 2005; Pushkaran AC et al. 2015).

LYZ is found in many human secretions such as tears, milk, mucus and saliva (Surna A et al. 2009; Minami J et al. 2015; Sahin O et al. 2016; Masschalck B & Michiels CW. 2003).

• An antibacterial role where the enzyme catalyzes the calcium-dependent hydrolysis of phospholipids in the bacterial membranes causing leakage of cell content (Buckland AG et al. 2000a,b; Foreman-Wykert AK et al. 1999; Beers SA et al. 2002; Nevalainen TJ et al. 2008)
• A proposed non-catalytic role in which PLA2G2A forms supramolecular aggregates with anionic phospholipid vesicles or debris (Bezzine S et al. 2002; Birts CN et al. 2008). These aggregates are then internalized via interactions with cell surface heparin sulphate proteoglycans and macropinocytosis for disposal by macrophages.

PLA2G2A shows bactericidal activity against various bacterial pathogens, in particular, against gram-positive bacteria such as Micrococcus luteus, Listeria innocua and Staphylococcus aureus though the sensitivity varies greatly between species (Buckland AG et al. 2000b; Beers SA et al. 2002; Koprivnjak T et al. 2002).

RNase A family is a vertebrate-specific gene family (Goo SM & Cho S 2013). Members of RNase A family share specific elements of sequence homology, a unique disulfide-bonded tertiary structure, and the ability to hydrolyze polymeric RNA (Beintema JJ & Kleineidam RG 1998; Rosenberg HF 2008). Eight catalytically active members are found in humans: RNase1 (pancreatic RNase), RNase2 (eosinophil derived neurotoxin/EDN), RNase3 (eosinophil cationic protein/ECP), RNase4, RNase5 (angiogenin), RNase6, RNase7 (skin-derived RNase), and RNase8 (divergent paralog of RNase7) (Sorrentino S 2010). Analysis of human genome sequence has revealed the existence of five additional RNases named as RNases 9-13, although they appear to lose enzymatic activity (Devor EJ et al. 2004; Castella S et al. 2004; Cho S et al. 2005). All human RNase A family members encode relatively small polypeptides of 14 to 16kDa containing signal peptides of 20 to 28 amino acids for protein secretion. Mature RNases contain 6 to 8 cysteine residues that are crucial to hold the overall tertiary structure (Sorrentino S 2010). Apart from the ribonuclease activity the RNase A family members have been implicated in a wide variety of biological actions including antipathogen and immunomodulatory activities (Harder J & Schroder JM 2002; Rudolph B et al. 2006; Boix E et al. 2008; Boix and Nogués, 2007; Spencer JD et al. 2011; Becknell B et al. 2015; Rosenberg HF 2015). Evidence of antimicrobial properties displayed by distantly related members ascribed to the family an ancestral role in host defence (Pizzo E & D’Alessio G 2007; Rosenberg HF et al. 2008).

RNase3, RNase6 and RNase7 have been identified as the most potent human antibacterial ribonucleases with a broad antimicrobial action against Gram-positive and Gram-negative bacteria (Pulido D et al. 2013, 2016; Zhang J et al. 2003; Boix E et al. 2008; Torrent M et al. 2010). Mutagenesis analysis revealed that ribonuclease-inactive RNase7 protein exhibited similar anti-microbial activity against P. aeruginosa, E. faecium and E. coli as the wild-type protein suggesting that RNase7 may kill bacteria independently of its ribonuclease catalytic activity (Huang YC et al. 2007; Koten B et al. 2009). Similar results were reported on microbicidal effect of ribonuclease-inactive RNase3 and 6 proteins against S. aureus (Rosenberg HF 1995; Pulido D et al. 2016a). Being cationic proteins with a high pI, RNase3, 6 and 7 interact with anionic components of biological membranes (Zhang J et al. 2003; Boix E et al. 2008; Torrent M et al. 2010; Boix E et al. 2012; Pulido D et al. 2016a). RNase3, 6 and 7 present, respectively, a high number of either Arg, His or Lys surface-exposed residues that may contribute to their distinct bactericidal mechanisms of action (Torrent M et al. 2010; Prats-Ejarque G et al. 2016). RNase3 displays a membrane disruption capacity that is dependent on both surface exposed hydrophobic and cationic residues. RNase3 can bind and partially insert into the lipid bilayers, promoting its aggregation and final lysis, following a carpet-like mechanism. The RNase3 agglutination process precedes the bacterial death and lysis event. The antimicrobial properties of the RNase6 are comparable to its RNase3 homolog and correlate to the bacterial cell damage and agglutination activities (Pulido D et al. 2016a). In contrast, RNase7 has no significant membrane aggregation capacity (Torrent M et al. 2010). RNase7 binds and permeabilizes the bacterial membrane displaying a much higher leakage capacity compared to RNase3 (Torrent M et al. 2010; Huang YC et al. 2007). Membrane permeabilization by RNase7 required four clustered lysine residues but no catalytic residues (Huang YC et al. 2007). Binding to PGN and LPS has been reported for RNases 3 and 7 (Torrent M et al. 2010; Pulido D et al. 2016b). Studies using a battery of progressively truncated LPS-defective E. coli strains correlated the LPS interaction with the protein cell agglutination and bactericidal activities (Pulido D et al. 2012). Further work indicated that RNase3 and RNase 6 high cell agglutination activity towards Gram negative species is retained by their respective N-terminus peptides (Torrent M et al. 2012, 2013; Pulido D et al. 2016c). In particular, the RNase3 N-terminus encompasses a specific patch (Y33-R36) required for LPS binding and an hydrophobic aggregation prone region (A8-I16) that mediates the protein self amyloid- like aggregation and promotes the cell death.