Subtilisin‐like proteases in plant defence: the past, the present and beyond

Summary

Subtilisin‐like proteases (or subtilases) are a very diverse family of serine peptidases present in many organisms, but mostly in plants. With a broad spectrum of biological functions, ranging from protein turnover and plant development to interactions with the environment, subtilases have been gaining increasing attention with regard to their involvement in plant defence responses against the most diverse pathogens. Over the last 5 years, the number of published studies associating plant subtilases with pathogen resistance and plant immunity has increased tremendously. In addition, the observation of subtilases and serine protease inhibitors secreted by pathogens has also gained prominence. In this review, we focus on the active participation of subtilases in the interactions established by plants with the environment, highlighting their role in plant–pathogen communication.

Subtilisin‐Like Proteases: History, Classification and General Features

In 1947, Lindstrom‐Lang and Ottesen accidentally discovered an extracellular alkaline proteinase during a study of the conversion of ovalbumin to plakalbumin (Lindstrom‐Lang and Ottesen, 1947). In this study, the authors used a dry powder of a bacterial enzyme from Bacillus subtilis and named it subtilisin, as homage to the organism in which it was observed for the first time (Lindstrom‐Lang and Ottesen, 1947). Between 1952 and 1954, Güntelberg's work in the Carlsberg laboratory led to the isolation and crystallization of this enzyme, named subtilisin Carlsberg (Güntelberg, 1954; Güntelberg and Ottesen, 1952, 1954). Four years later, another subtilisin, BPN′ (a subtilisin from Bacillus subtilis strain N′), was also isolated and crystallized (Hagihara et al., 1958). The physical properties and amino acid composition of this BPN′ subtilase were first described in 1965 by Matsubara and co‐workers (Matsubara et al., 1965). The complete amino acid sequences of the subtilisins Carlsberg and BPN′ were only revealed in 1966 (Smith et al., 1966) and the structure of the subtilisin BPN′ was solved at 2.5‐Å resolution in 1969 (Wright et al., 1969). In plants, the first purified subtilisin was the cucumisin from sarcocarp of melon fruit in 1975 (Kaneda and Tominaga, 1975). However, only two decades later, the complete amino acid sequence of this protein was described (Yamagata et al., 1994). Other plant subtilases were characterized throughout the years, but only in 2004 were subtilases associated with plant cell death events with the identification of a partial sequence of two subtilases exhibiting caspase activity: SAS‐1 and SAS‐2 (Coffeen and Wolpert, 2004).

In 1996, the MEROPS database (http://merops.sanger.ac.uk; Rawlings and Barrett, 1999) was created and a hierarchical classification for peptidases was established. This classification involved the clustering of homologous sets of peptidase and protein inhibitor sequences into peptidase and inhibitor species, which were, in turn, clustered into families and grouped into clans (Rawlings and Barrett, 1999). Subtilisin‐like proteases (also known as subtilases) were classified as members of the S8A subfamily within the second largest family of serine peptidases (family S8), belonging to the SB clan (Rawlings and Barrett, 1993, 1994). Subtilases are found in archaea, bacteria, eukarya, fungi and yeasts (Siezen et al., 1991), being highly represented in plants.

Since the discovery of the first subtilisin‐like protease, several subtilases have been identified and characterized in the most diverse organisms, not only individual proteins but also complete subtilase gene families. The first subtilase gene family to be described belonged to Lycopersicon esculentum (tomato) and consisted of 15 members (Meichtry et al., 1999). After this work, subtilase gene families were also characterized in Arabidopsis thaliana, presenting 56 known genes (Rautengarten et al., 2005), in Oryza sativa (rice) with 63 genes (Tripathi and Sowdhamini, 2006), in moss Physcomitrella patens with 23 genes, in Populus trichocarpa with 90 genes (Schaller et al., 2012) and, more recently, in Vitis vinifera and Solanum tuberosum (potato), both with 82 known genes (Figueiredo et al., 2016; Norero et al., 2016).

Plant subtilases present a broad spectrum of biological functions, being involved not only in all aspects of the plant life cycle, such as the development of seeds and fruits, cell wall modification, processing of peptide growth factors and epidermal development, but also in the response to biotic and abiotic stress (reviewed in Schaller et al., 2012).

Major milestones concerning subtilase history and relation to pathogen resistance are presented in Fig. 1.

Figure 1.

Major milestones in the history of subtilases, emphasizing the discoveries associated with plant–pathogen interactions. CEV, citrus exocortis viroid; JA, jasmonic acid; MeJA, methyl jasmonate; PCD, programmed cell death; RuBPCase, ribulose bisphosphate carboxylase; SA, salicylic acid; TMV, Tobacco mosaic virus.

Structure and Biochemical Characteristics of Plant subtilases

Most of the plant subtilases are synthesized as inactive pre‐pro‐protein precursors whose structure is composed of three conserved domains. The catalytic domain, named the peptidase S8 domain, is present in all the subtilases and consists of a highly conserved catalytic triad composed of aspartate (Asp), histidine (His) and serine (Ser) amino acid residues (Dodson and Wlodawer, 1998) (Fig. 2). Some subtilases may also present a conserved catalytic asparagine (Asn) residue within this domain (Dodson and Wlodawer, 1998; Jordá et al., 1999; Siezen and Leunissen, 1997). The majority of subtilases also present a protease‐associated (PA) domain consisting of an insertion of 120–160 amino acids between the His and Ser active site residues (Luo and Hofmann, 2001). This PA domain is responsible for the displacement of the reactive Ser from the catalytic triad to the C‐terminus (Siezen and Leunissen, 1997) (Fig. 2). In many subtilases, the PA domain is also responsible for the homodimerization of the protein in order to activate it (Ottmann et al., 2009; Rose et al., 2010; Siezen and Leunissen, 1997). The PA domain has also been implicated in protein–protein interactions and substrate recognition (Rautengarten et al., 2005; Schaller et al., 2012). Studies in soybean have shown that this domain plays an important role in substrate selection, namely in substrate length determination (Rautengarten et al., 2005; Tan‐Wilson et al., 2012). There is one last conserved domain, also present in the majority of plant subtilases, named the inhibitor I9 domain or prodomain, which works as an auto‐inhibitory domain maintaining the inactive state of the zymogen and preventing the access of the substrate to the active site. This domain also works as an intramolecular chaperone that is transiently required to assist in catalytic domain folding (Baker et al., 1993; Bryan, 2002; Huang et al., 1997; Li and Inouye, 1994; Zhu et al., 1989).

Figure 2.

Domain architecture of SISBt3 (PDB ID: 3I6S; UniProtKB AC: O82777), a subtilase purified from Solanum lycopersicum (tomato) cell culture, showing the characteristic domains [signal peptide (Met¹–Ala²²), prodomain (Thr²³–Glu¹¹⁰), catalytic subtilase domain (Thr¹¹³–Arg³⁴² and Leu⁴⁷³–Tyr⁶⁵⁴), protease‐associated (PA) domain (Thr³⁴³–Tyr⁴⁷²) and C‐terminus fibronectin (Fn)‐III domain (Pro⁶⁵⁵–Trp⁷⁶¹)], as well as the three amino acid residues [aspartate (D¹⁴⁴), histidine (H²¹⁵) and serine (S⁵³⁸)] in the catalytic subtilase domain (Ottmann et al., 2009). This plant subtilase is the only one for which the domain architecture has been elucidated experimentally. Ala, alanine; Arg, arginine; Glu, glutamate; Leu, leucine; Met, methionine; Pro, proline; Thr, threonine; Trp, tryptophan; Tyr, tyrosine.

Interaction between the PA and inhibitor I9 domains leads to the cleavage of the N‐terminus, allowing the access of the substrates to the catalytic site, promoting subtilase activity (Bergeron et al., 2000). Most plant subtilases also present a signal peptide which targets the proteases for secretion. The removal of this peptide is a prerequisite for enzyme maturation and occurs in the endoplasmic reticulum (ER) or in the Golgi complex, either as an intramolecular autocatalytic reaction or as a result of the interaction with a secondary peptidase (Bergeron et al., 2000; Cedzich et al., 2009; Chichkova et al., 2010). In A. thaliana, an additional domain was also found, the fibronectin (Fn)‐III‐like domain, which is required for the activity of some of these enzymes, but dispensable in others (Rawlings and Salvesen, 2013).

Another feature of most plant subtilases is the apparent calcium (Ca²⁺) independence, contrary to that expected from modelling studies in the subtilisin BPN′ (Alexander et al., 2001; Siezen and Leunissen, 1997). In 2009, Ottmann and co‐workers demonstrated that the thermostability and activity of the subtilase SISBT3 from Solanum lycopersicum were not influenced by the addition of Ca²⁺ or chelating agents (Ottmann et al., 2009). Instead, in SISBT3, a positively charged site chain of Lys⁴⁹⁸ mimics the calcium ion bound function. This region, including the stabilizing lysine (Lys) residue, is highly conserved in all the plant subtilases studied so far (Rose et al., 2010). However, there are plant subtilases that depend on Ca²⁺ for activity. An example of a calcium‐dependent subtilase is ARA12 from A. thaliana, whose proteolytic activity is stimulated by the presence of Ca²⁺ ions (Hamilton et al., 2003). Indeed, three‐dimensional structural studies revealed the presence of Ca²⁺ ion‐binding pockets in this protein (Smith et al., 1999). The binding of calcium presents a stabilizing effect on this subtilase activity by reducing molecular flexibility, which is thought to be important in preventing thermal denaturation and autolysis (Siezen et al., 1991).

Concerning the impact of inhibitors in subtilase activity, some subtilases, such as A. thaliana ARA12, are easily inhibited by classical serine protease inhibitors (Hamilton et al., 2003). In contrast, other subtilases are inhibited by diisopropyl fluorophosphate (DFP), but unaffected by reducing compounds, such as β‐mercaptoethanol, like the cucumisins from melon fruit (Kaneda and Tominaga, 1975). The wheat subtilase EP3 is completely inhibited by 4‐(2‐aminoethyl)benzenesulfonyl fluoride (AEBSF) hydrochloride and phenylmethanesulfonyl fluoride (PMSF), less inhibited by ethyleneglycol‐bis‐(2‐amino ether)tetraacetic acid (EGTA) and not inhibited by other metalloprotease inhibitors, such as ethylenediaminetetraacetic acid (EDTA) and 1,10‐phenanthroline (Wang et al., 2013). In addition, the EP3 protease is insensitive to the serine/cysteine protease inhibitors leupeptin and N‐p‐toluenesulfonylbenzene propionyl chloride methyl ketone (TPCK) (Wang et al., 2013).

One important characteristic in plant subtilases is glycosylation, a post‐translational modification that regulates subtilase activity (Bykova et al., 2006; Cedzich et al., 2009). The most important protein glycosylation form is N‐linked, formed by the covalent attachment of Asn‐linked carbohydrates to the protein (Bykova et al., 2006; Gupta and Brunak, 2002). Protein N‐glycosylation has been described previously in P69B from tomato, being confirmed by mass spectrometry, with four N‐glycosylation sites with five glycoforms for each site in this subtilase (Bykova et al., 2006).

Subtilases Involved in Plant Defence

Subtilases have been implicated in several plant development functions, but have gained more prominence as a result of their involvement in plant responses to both abiotic and biotic environmental stimuli.

Following an abiotic stimulus, subtilases have been reported to be involved in drought and salt resistance mechanisms (Budič et al., 2013; Che et al., 2010; Liu et al., 2007; Liu and Howell, 2010a, 2010b). A well‐characterized example is the A. thaliana subtilase AtSBT6.1 associated with the unfolded protein response on salt stress through the cleavage of an ER‐resident type II membrane protein (bZIP28). The cleavage of the bZIP28 protein is essential for the activation of genes associated with the salt stress response (Fig. 3A).

Figure 3.

Illustration of subtilase participation in plant defence mechanisms after biotic or abiotic stress (A–I). Plant subtilases are represented in blue and pathogen subtilases in red. CEV, citrus exocortis viroid; ER, endoplasmic reticulum; HG, homogalacturonan; JA, jasmonic acid; LA, linolenic acid; LRP, leucine‐rich repeat protein; PA, phytodienoic acid; PCD, programmed cell death; PHB, prohibitin; PME, pectin methylesterase; PR, pathogenesis‐related; SBT, subtilase.

Reports of subtilase participation in resistance responses following biotic stimuli have increased over recent years. The first evidence of subtilase involvement in plant–pathogen interactions was reported in tomato plants by Granell et al. (1987). These authors identified the accumulation of the subtilase P69 in tomato leaves after infection with citrus exocortis viroid (CEV). This subtilase was characterized as an alkaline proteinase located in the vacuole and in the intercellular spaces of leaf parenchyma cells. It was associated with engulfed ‘inclusion bodies’ and dispersed material, and with the degradation of the large subunit of RuBPCase (ribulose bisphosphate carboxylase, the most abundant protein of the chloroplast). It was hypothesized that this tomato P69 acts during the last stages of chloroplast degradation when contents are released to the cytosol (Vera et al., 1989; Vera and Conejero, 1988). In 1996, the P69 subtilase was cloned by Tornero et al. (1996a). Later, it was shown that the P69 subtilase consisted of a gene family with six closely related members, named P69A to P69F (Jordá et al., 1999, 2000). These P69 genes were shown to be expressed in different conditions. P69A, P69D and P69E were associated with developmental processes, whereas P69B and P69C were shown to behave as pathogenesis‐related (PR) genes, as both were transcriptionally activated on infection with Pseudomonas syringae (Jordá et al., 1999, 2000). It was also shown that P69B and P69C activation was not restricted to the point of pathogen inoculation, but was extended throughout the afflicted leaf blade, suggesting a long‐distance signalling process. Experiments with salicylic acid (SA), a signal molecule that mediates the long‐distance activation of the plant defence reaction, revealed that both P69B and P69C were also up‐regulated by this immunity elicitor (Jordá et al., 1999; Tornero et al., 1997).

P69 was also the first plant subtilase for which protein substrates were identified, such as systemin (Schaller and Ryan, 1994) and the leucine‐rich repeat protein (LRP) (Tornero et al., 1996b). Systemin is an 18‐amino‐acid polypeptide derived from the larger precursor prosystemin by proteolytic processing in secretory vesicles by members of a family of site‐specific, subtilisin‐related proteinases (Bergey et al., 1996). It plays a fundamental role in response to herbivore damage or mechanical wounding through the systemic signalling pathway activation of plant defence genes involved in the reprogramming of the protein synthesis machinery. Systemin also participates in a lipid‐based signalling cascade through the activation of a lipase in receptor cell membranes, resulting in the release of linolenic acid, the jasmonic acid (JA) precursor (Bergey et al., 1996). This pathway activation leads to the synthesis of systemic wound response proteins (SWRPs), which include serine proteinase inhibitors (Bergey et al., 1996) (Fig. 3B). The other identified substrate, LRP, mediates molecular recognition and/or interaction processes in the extracellular matrix of eukaryotic cells under normal and/or PR conditions to initiate different signalling processes (Tornero et al., 1996b). In the tomato–viroid infection context, it was shown that tomato plants activate a set of genes encoding PR proteins, such as the subtilase P69, which is responsible for the proteolytic processing of the LRP (Tornero et al., 1996b). Later, it was discovered that these two P69 substrates are related, and now it is known that systemin is perceived at the cell surface by a leucine‐rich repeat receptor‐like kinase that induces the JA synthesis pathway at the site of wounding as a prerequisite for systemic defence gene induction (Howe and Schaller, 2008; Ryan, 2000; Wasternack et al., 2006) (Fig. 3B).

After the discovery of the P69 involvement in tomato resistance against CEV and P. syringae, the protein isoform P69B was also associated with the defence response against Phytophthora infestans (Tian, 2005; Tian et al., 2004). At the same time, it was also found that P. infestans secretes serine protease inhibitors of the Kazal family, including the two‐domain EPI1 protein, that inhibit and interact with the P69B subtilase of tomato (Tian, 2005; Tian et al., 2004) (Fig. 3C). These results suggest that there is a concerted interaction between the host subtilase P69B and the pathogen serine protease inhibitors, and that this interaction dictates the success of pathogen colonization (Tian, 2005; Tian et al., 2004). A year earlier, Zhao et al. (2003) highlighted the involvement of P69B and JA/wound response genes for a successful defence response to P. syringae pv. tomato strain DC3000. Later, also in tomato, it was shown that, after inoculation with Ralstonia solanacearum, a stem cell wall subtilase protein was highly accumulated in the resistant genotype (Hawaii7996) (Dahal et al., 2010).

In grapevine, subtilases were identified in response to two biotrophic pathogens, Plasmopara viticola and Erysiphe necator, the causing agents of downy and powdery mildews, respectively. In the V. vinifera–P. viticola pathosystem, several studies have highlighted the involvement of subtilases in the establishment of a successful resistance response (Figueiredo et al., 2008, 2012; Monteiro et al., 2013). When comparing resistant and susceptible grapevine genotypes, a subtilase presenting sequence similarity with tomato P69C was observed to be constitutively expressed in the resistant genotypes, increasing its expression after inoculation (Figueiredo et al., 2008, 2012; Monteiro et al., 2013). More recently, subtilase participation in the defence response against P. viticola was reinforced (Figueiredo et al., 2016). In Vitis pseudoreticulata leaves infected with E. necator, an up‐regulation of several subtilase genes was also reported (Weng et al., 2014).

In 2013, studies in A. thaliana identified a serine protease coding gene, SBT3.3, whose protein accumulates in the extracellular matrix. The authors hypothesized that this subtilase may cleave the extracellular domain (ectodomain) of a larger protein acting as a receptor located in the plasma membrane (Ramírez et al., 2013). Thus, as a consequence of the proteolytic shedding of the ectodomain, the receptor could become activated and initiate a downstream immune signalling process. Thereafter, a positive feedback loop circuit would maintain SBT3.3 expression at a level sufficient to keep cells in a sustained sensitized mode. This expression pattern would consequently be the basis to explain the memory‐based characteristics of priming and induced resistance (Ramírez et al., 2013). Considering the high sequence similarity of this enzyme with tomato P69C, the authors suggested that its substrate may also be an extracellular LRP. Moreover, Ramírez et al. (2013) have shown that the expression of SBT3.3 rapidly increases during the activation of innate immunity preceding the activation of SA‐responsive genes.

In coffee leaves, a proteomic analysis of apoplastic fluid in incompatible (resistant) and compatible (susceptible) Coffea arabica–Hemileia vastatrix interactions has also revealed an increase in subtilase proteins at all time points in the resistant genotype (Guerra‐Guimarães et al., 2015).

In the past year, several studies have highlighted the involvement of subtilases in plant resistance to pathogens. In cotton, a subtilase called GbSBT1 has been identified, characterized and related to Verticillium dahliae‐induced resistance (Duan et al., 2016). In this study, the authors observed that GbSBT1 knockdown reduces cotton defence against V. dahliae attack. This subtilase is localized in the cell membrane and in the extracellular apoplast, where the plant–pathogen interaction occurs and which serves as the recognition site between cotton and V. dahliae. The authors also found that this subtilase interacts with a prohibitin (PHB)‐like protein expressed by V. dahliae during infection, which is associated with ethylene‐ or nitric oxide‐mediated cellular responses in plants (Duan et al., 2016). It was hypothesized that the PHB from V. dahliae interacts with cotton GbSBT1 and activates the transcription of genes related to Verticillium resistance signalling (Fig. 3D). Moreover, when cotton plants were treated with JA and ethylene, the GbSBT1 gene was activated and the protein moved to the cytoplasm, demonstrating that JA signalling is required for plant resistance against V. dahliae (Fig. 3D). Interestingly, these authors also discovered that the ectopically expressed GbSBT1 gene enhances the resistance of Arabidopsis to Fusarium oxysporum and to V. dahliae infections, and activates the expression levels of defence‐related genes (Duan et al., 2016). In wheat, a subtilase has been identified and partially characterized in the apoplastic space (Fan et al., 2016). After Puccinia triticina infection, which causes leaf rust, an increase in protease activity was observed during the incompatible interaction, which was connected to this subtilase. This activity increased until a late stage of wheat leaf development and increased in response to both heat shock and biotic stress (Fan et al., 2016). In S. tuberosum, several subtilase genes have been shown to increase their expression after P. infestans infection (Norero et al., 2016). The involvement of plant subtilases in herbivore resistance has also been reported for the first time. Meyer et al. (2016) highlighted the involvement of a previously described subtilase, SISTB3 (Cedzich et al., 2009), in tomato resistance against Manduca sexta larvae. These authors found an enhanced performance of M. sexta larvae on SISBT3‐silenced plants, suggesting that SISBT3 activity may be required locally in the late stages of induced herbivore defence in tomato plants. In addition, considering the apparent co‐localization in the tomato vasculature of the subtilase SISBT3, the prosystemin (the large precursor of the peptide hormone systemin) and the enzymes involved in JA biosynthesis, it was hypothesized that SISBT3 is involved in systemin processing and the JA‐mediated resistance response (Meyer et al., 2016). Moreover, there were observed changes in cell wall composition in transgenic plants with altered SISBT3 expression levels, suggesting a potential involvement of this subtilase in the control of pectin methylesterase (PME) activity (Senechal et al., 2014). PME catalyses the de‐methylesterification of homogalacturonan (HG), the major pectin constituent of the primary cell wall (Senechal et al., 2014), with dramatic consequences on the mechanical properties and digestibility of the cell wall (Peaucelle et al., 2011), affecting the attack performance of herbivorous insects (Calderón‐Cortés et al., 2012; Körner et al., 2009) (Fig. 3E). Furthermore, in this reaction, substantial amounts of methanol are released, and it was hypothesized that the released methanol may be directly toxic to larvae and/or may serve as a signal for direct and indirect plant defence responses (Meyer et al., 2016) (Fig. 3E). Finally, considering the SISBT3 stability and its high proteolytic activity at alkaline pH, it was shown that SISBT3 remained intact in M. sexta midgut, suggesting that it may be involved in the degradation or processing of proteins in the insect digestive system (Meyer et al., 2016) (Fig. 3E).

Very recently, it has been reported that the A. thaliana subtilase AtSBT6.1 is responsible for the processing of the pro‐peptide form of RAPID ALKALINIZATION FACTOR 23 (RALF23) (Stegmann et al., 2017). After cleavage, the released RALF23 binds to malectin‐like receptor kinase FERONIA (FER). FER is present in the plasma membrane and acts as a scaffold to regulate immune receptor complex formation. Contrary to the previous observations for plant subtilases, it was shown that the binding of RALF23 to FER promotes a negative feedback mechanism that inhibits the scaffolding function of FER and dampens immune signalling on pathogen attack. The authors suggested that fungal pathogens secrete RALF23 homologues that suppress plant immunity by inhibition of the formation of active receptor complexes. This study suggests that subtilases, like AtSBT6.1, can act as negative regulators of immunity (Stegmann et al., 2017).

Subtilases in Plant Programmed Cell Death

Cell death plays a central role in innate immune responses in both plants and animals (Coll et al., 2011). In plants, pathogen recognition leads to the inhibition of pathogen growth, which is often accompanied by the triggering of the hypersensitive response (HR), a form of programmed cell death (PCD) localized at the site of attempted pathogen invasion.

Current data indicate that some specific subtilases play a role in plant PCD (Chichkova et al., 2010; Coffeen and Wolpert, 2004; Fernández et al., 2012, 2015). These proteases, displaying amino acid sequence similarity to subtilases and containing an active‐site Ser residue, exhibit caspase specificity (Chichkova et al., 2010; Coffeen and Wolpert, 2004). This specific group of subtilases can be divided into two subgroups, saspases and phytaspases, that are distinguished by their substrate specificity and role in PCD [reviewed in Kabbage et al. (2017) and Vartapetian et al. (2011)]. Saspases are extracellular proteins responsible for RuBisCO (ribulose‐1,5‐bisphosphate carboxylase/oxygenase) proteolysis in PCD, and prefer the peptide recognition pattern of caspases, such as IETD and LEHD. However, phytaspases are extracellular proteins in healthy tissues that are shifted to cytoplasm on PCD induction. In PCD, phytaspases are responsible for the cleavage of foreign proteins upstream of mitochondrial events. Considering their caspase specificity, phytaspases have preference for peptide recognition patterns such as VEID, IETD, LEHD and VDVAD [reviewed in Kabbage et al. (2017) and Vartapetian et al. (2011)].

In 2004, two proteases characterized as subtilases that exhibited caspase specificity were identified in Avena sativa and named SAS‐1 and SAS‐2 (saspases) (Coffeen and Wolpert, 2004). It was shown that these saspases participated in a PCD‐induced protease cascade leading to RuBisCO cleavage. These saspases were constitutively present in an active form and re‐localized to the extracellular fluid after induction of PCD by either victorin, a toxin produced by the fungus Cochliobolus victoriae, or heat shock (Coffeen and Wolpert, 2004). However, the function of saspases in PCD execution remains unknown.

Also in 2004, Chichkova and co‐workers identified a protein in tobacco with caspase‐like activity (Chichkova et al., 2004) which, in 2010, was recognized as a subtilase with caspase specificity, named phytaspase, when, after mutation of the predicted catalytic S⁵³⁷ residue, the enzyme lost its proteolytic activity (Chichkova et al., 2010). In tobacco plants, this enzyme was showed to be inactive in healthy tissues, but was activated in Tobacco mosaic virus (TMV)‐induced HR. Moreover, in tobacco infected with Agrobacterium tumefaciens, this phytaspase displayed an exceptional selectivity, introducing a single break in the A. tumefaciens VirD2 protein, after the D⁴⁰⁰ residue within the TATD motif. This cleavage resulted in the detachment of the C‐terminus nuclear localization signal (NLS)‐containing peptide of VirD2, which is essential for nuclear uptake of foreign DNA within the plant cell during bacterial infection and plant transformation (Shurvinton et al., 1992; Steck et al., 1990) (Fig. 3F). Indeed, VirD2 inactivation by tobacco phytaspase may represent a protective mechanism aimed at limiting the delivery and expression of foreign genes in plants (Chichkova et al., 2004). The authors showed that phytaspase is constitutively secreted into the apoplast before PCD, but, unexpectedly, is partially re‐imported into the cell during PCD (Chichkova et al., 2004).

More recently, a subtilase with caspase‐3 activity, named StSBTc‐3, has been identified and characterized in the S. tuberosum–P. infestans interaction, distinct from the saspases and phytaspases already described (Fernández et al., 2012, 2015). The authors showed that StSBTc‐3 is the major protein in the apoplast of detached potato leaves after infection and that this enzyme shows DEVDase activity (preferred DEVD peptide recognition pattern) (Fernández et al., 2012). In addition, StSBTc‐3 can produce in vitro cytoplasm shrinkage in plant cells and plant cell death, possibly through the completion of HR‐like necrotic reactions that restrict pathogen spread in potato leaves (Fernández et al., 2012). Like tobacco phytaspases, StSBTc‐3 activity in potato leaf apoplast is constitutive in healthy tissues, contrary to saspases, whose activity in oat extracellular fluids is only detected after the induction of PCD. However, unlike phytaspases which, in response to death‐inducing stimuli, are re‐localized to the cell interior, potato StSBTc‐3, like saspases, is induced in the potato leaf apoplast on pathogen infection. Moreover, neither phytaspases nor saspases could hydrolyse a DEVD‐based caspase substrate, and so it is suggested that StSBTc‐3 belongs to a new subgroup of subtilases with caspase specificity, in this case, the subgroup of serine peptidases with DEVDase activity (Fernández et al., 2012).

Studies with serine peptidase inhibitors have shown that common subtilases, without caspase specificity, are also involved in PCD. In the V. vinifera–P. viticola interaction, studies using resistant and susceptible grapevine genotypes have shown that the use of serine inhibitors partially inhibits the overall activation of PCD and thereby changes the level of susceptibility of grapevine towards P. viticola (Gindro et al., 2012). Indeed, after plant treatment with these inhibitors, an increase in the P. viticola infection rate was observed in all the grapevine genotypes, overcoming the plant defence barriers and completing its life cycle. This result also reinforces the involvement of subtilase in grapevine defence mechanisms. The authors proposed that the secretome of P. viticola may be specific, but may also be tailored to the host plant to a certain extent. It was hypothesized that some components of the P. viticola secretome could inhibit the endogenous subtilases of susceptible varieties, thereby inhibiting the plant's normal defence reaction, whereas resistant or immune varieties may possess endogenous subtilases that are not recognized because of slight structural modifications of the protein patterns of these cultivars. In this case, plant defence mechanisms would continue to operate, with fatal consequences for the pathogen and restricting its development (Gindro et al., 2012).

More recently, in Arabidopsis, the subtilase AtSBT5.2 was unexpectedly associated with MYB30 (Serrano et al., 2016), a transcription factor that promotes defence and cell death‐associated responses through the transcriptional activation of genes related to the lipid biosynthesis pathway (Raffaele et al., 2008). Serrano et al. (2016) have also shown that the AtSBT5.2 transcript presents two splice variants. The AtSBT5.2(b) variant encodes an atypical subtilase that specifically mediates the retention of MYB30 at the endosomal vesicles, stopping MYB30 from entering the nucleus, after bacterial treatment. Thus, MYB30 cannot bind to the DNA or activate its target genes, weakening the defensive response mediated by this transcription factor (Fig. 3G). Thus, AtSBT5.2(b) is an additional negative regulator of MYB30 activity (Serrano et al., 2016) and, unlike most subtilases, appears to play a role in plant infection.

Elicitors and Small Peptides in Plant–Pathogen Interactions: The Role of Subtilases

In plants, elicitors are widely used to induce and study defence response mechanisms that mimic the response on pathogen attack. Subtilases have been shown to increase their expression in plants after elicitor treatment. In Sorghum bicolor, a subtilase has been shown to increase its expression after elicitation with methyl jasmonate (MeJA), SA and aminocyclopropane carboxylic acid (ACC) (Salzman et al., 2005). Although SA is thought to block JA synthesis (Pena‐Cortés et al., 1993), the authors observed an SA‐ and MeJA‐mediated induction of putative homologues of genes composing the entire octadecanoid (ODA) pathway for JA synthesis (Salzman et al., 2005), this pathway being connected to the plant defence response. Moreover, SA/JA co‐regulation has been reported to be associated with resistance against some pathogens (Berrocal‐Lobo et al., 2002). Thus, a possible link between subtilases and both SA and JA pathways should not be discarded.

In V. vinifera treated with methylated cyclodextrins (MBCD), a significant induction of a subtilase involved in the release of mucilage in seeds has been reported, suggesting a possible participation of this protein in the building of physical barriers to pathogen invasion (Martinez‐Esteso et al., 2009).

Several studies have shown that plants themselves synthesize small peptides similar to pathogen‐derived elicitors. These small peptides trigger or amplify the plant innate immune response (Pearce et al., 2010). In soybean, one of these small peptides has been characterized and identified as part of a subtilase. On pathogen attack or wounding, the conserved PA domain of the extracellular subtilase Glyma18g48580 is cleaved into a 12‐amino‐acid peptide (GmSubPep: NTPPRRAKSRPH) which acts as an endogenous elicitor and initiates defence signalling pathways (Pearce et al., 2010; Yamaguchi et al., 2011). It has been suggested that, on exposure to a stimulus, this endogenous peptide would be available for receptor binding, initiation of defence signalling pathways and activation of defence‐related genes (Pearce et al., 2010). Indeed, the treatment of soybean cells with this peptide increased the expression of cytochrome P450 gene (Cyp93A1), involved in the synthesis of phytoalexin (an antimicrobial and antioxidative metabolite synthesized by plants which accumulates rapidly in areas of pathogen infection) (Ahuja et al., 2012), and the expression of the PR chitinase 1b‐1 gene (Chib‐1b) (Pearce et al., 2010).

Subtilases Involved in Pathogenicity

It is evident that plant subtilases play important roles in defence responses. In recent decades, several reports have also shown that plant invaders also present subtilases that might be involved in the mechanism of pathogen attack. The presence of serine proteases in the secretome of various pathogens, in infection situations, has long been observed (reviewed in Xia, 2004).

In 1998, Olivieri's group observed the accumulation of a secreted subtilase in potato tubers after Fusarium eumartii infection (Olivieri et al., 1998). Fusarium eumartii is a phytopathogenic fungus that causes potato dry rot, invading the wounded potato parenchyma cells, and, like most fungi, secretes a broad spectrum of proteolytic enzymes during the penetration and colonization of plant tissue. A fungal subtilase that degraded potato PR proteins as well as specific polypeptides of intercellular washing fluids and cell walls was identified (Olivieri et al., 1998, 2002) (Fig. 3H). These findings led the authors to propose a possible contribution of this subtilase to the invasion strategy of F. eumartii (Olivieri et al., 2002).

In wheat Fusarium head blight (FHB) disease, caused by Fusarium graminearum (Gottwald et al., 2012), the pathogen uses a specific arsenal of virulence factors, including subtilases, which occur during almost the entire course of the disease (Pekkarinen and Jones, 2003; Walter et al., 2010). A gene expression profiling study in the wheat–F. graminearum interaction has shown that subtilases are released in infected wheat kernels mainly to disrupt host cell membranes during necrotrophic intracellular nutrition (Fig. 3I). However, at the same time, a differential up‐regulation of five serine protease inhibitors was also observed in wheat (Gottwald et al., 2012). Another study, in Triticum dicoccum (emmer) grains infected with F. graminearum and Fusarium culmorum, it has been shown that infection of Fusarium leads to the induction of the serine protease inhibitors serpin‐Z1A and serpin‐Z2B by emmer grains (Eggert et al., 2011). Nevertheless, the inhibition of fungal proteinases by the endogenous emmer protease inhibitor serpin prevents, to a certain degree, the digestion of seed storage proteins after fungal infection (Pekkarinen et al., 2007; Phalip et al., 2005). These findings suggest that subtilases and serine protease inhibitor proteins are involved in the crosstalk between pathogen and host. Indeed, during plant–pathogen interactions, it is expected that both organisms release specific sets of proteases and protease inhibitors mutually impairing each other (López‐Otín and Overall, 2002).

More recently, Chalfoun et al. (2013) purified and characterized an extracellular subtilase, AsES, which acts as a plant defence response elicitor, produced by an avirulent isolate of the strawberry pathogen Acremonium strictum. The results showed that AsES can trigger a strong defence reaction in strawberry plants, characterized primarily by a transient oxidative burst, followed by a strong transcriptional induction of PR‐1 (FaPR1), class II chitinase (FaChi2–1) and, finally, manifested by an enhanced resistance against fungal pathogens of hemibiotrophic (i.e. Colletotrichum acutatum) and necrotrophic (i.e. Botrytis cinerea) lifestyles. However, the authors concluded that the proteolytic activity is necessary, but not sufficient, to induce defence, and suggested that AsES might induce defence by means of proteolysis of one or multiple host proteins that are specific targets of this protease (Chalfoun et al., 2013).

Lastly, in the entomopathogenic fungus Verticillium lecanii, a biocontrol agent, the production of a subtilase, named VlPr1, important in the biological control activity of some insect pests by degrading insect cuticles, was observed (Yu et al., 2012). Indeed, during in vitro experiments, VlPr1 showed a broad spectrum of antifungal activity towards fungal pathogens that devastate plants, particularly Fusarium moniliforme, Penicillium italicum, Magnaporthe oryzae and Exserohilum turcicum (Yu et al., 2012). These results show a new role of subtilases in plant–pathogen interactions. In this case, the subtilase is produced by an organism living in symbiosis with the plant and its function is to help the plant to defend itself on pathogen attack.

Conclusions

Plant–pathogen communication is very complex with a myriad of processes being activated by both partners, leading to the establishment of compatible or incompatible interactions. In this review, we have highlighted the involvement of subtilases secreted by both interaction partners. Although plant subtilases seem to be involved, not only in mounting an effective defence strategy through the activation of signalling cascades, but also in causing direct damage to the pathogen, pathogens secrete subtilases in order to suppress plant defences. Despite all the recent advances in subtilases associated with plant defence mechanisms, very little is known about their structure, post‐translational modification sites, substrates and function. Moreover, knowledge on the cellular targets and cellular interacting partners of host proteases is still scarce. Advances in both transcriptomic and proteomic fields, as well as in structural methods, will contribute to an understanding of the significance of subtilases in the regulation of plant biotic stress responses. The activation of plant or pathogen serine proteases and their expected functional complexity suggest exciting new future discoveries in the field of regulatory proteases in plant–pathogen interactions.

Author Contributions

J.F., M.S.S. and A.F. conceived and wrote the manuscript.

Conflict of Interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as potential conflicts of interest.