Introduction

Feruloyl esterases

Feruloyl esterases [E.C. 3.1.1.73; also known as ferulic acid esterases (FAE), cinnamoyl esterases, cinnamic acid hydrolases] are a subclass of the carboxylic acid esterases (E.C. 3.1.1.1) that are able to hydrolyse the ester bond between hydroxycinnamic acids and sugars present in the plant cell walls (Williamson et al. 1998b). These enzymes have previously been classified as type A (e.g. Aspergillus niger FaeA) and type B (e.g. A. niger CinnAE, FaeB, Penicillium funiculosum FaeB, Neurospora crassa Fae-1), based on their specificity for the substrate aromatic moiety, specificity for the linkage to the primary sugar and ability to release ferulic acid dehydrodimers from esterified substrates (Crepin et al. 2003a; De Vries et al. 2002b; Kroon et al. 1999, 2000). The substrate utilisation of either methyl sinapate or methyl caffeate by feruloyl esterases is the basis of this classification. In addition, the majority of these esterases show acetyl xylan esterase activity, regardless of whether they are assigned as acetyl or feruloyl esterases. However, specific acetyl xylan esterase activity is rather low for the enzymes classified as feruloyl esterases. The two major feruloyl esterases of A. niger have received most attention, namely AnFaeA and AnFaeB (De Vries et al. 1997, 2002b; Faulds and Williamson 1994; Kroon et al. 1996, 1997).

Type A feruloyl esterases

Type A feruloyl esterases tend to be induced during growth on cereal-derived substrates. The enzyme releases ferulic acid from 1,5-ester-linked feruloylated arabinose (Ralet et al. 1994) and is also able to release low quantities of 5,5′- and 8-O-4′-ferulate dehydrodimers from plant material when pre-treated with a xylanase or when a xylanase is co-incubated with the feruloyl esterase (Bartolome et al.1997; Faulds and Williamson 1995; Kroon et al. 1999; Williamson et al. 1998a). There is no report of these enzymes releasing diferulate from pectin-derived material. Type A feruloyl esterases also show a preference for the phenolic moiety of the substrate that contains methoxy substitutions, especially at carbon(s) 3 and/or 5, as occurs in ferulic and sinapic acids. Regarding specificity against synthetic substrates, type A feruloyl esterases are active against methyl ferulate, methyl sinapate and methyl p-coumarate, but not methyl caffeate. They appear to prefer hydrophobic substrates with bulky substituents on the benzene ring (Kroon et al. 1997, 1999; Williamson et al. 1998a).

Type B feruloyl esterases

Type B feruloyl esterases are preferentially secreted by growth on sugar beet pulp. The enzymes release ferulic acid ester-linked to either C-2 of feruloylated arabinose or C-6 of feruloylated galactose residues (Kroon and Williamson 1996), but they are unable to release the dimeric forms of ferulate. The A. niger type B enzyme also works on 1,5-linked ferulic acid as found in cereals but at a lower rate than the type A esterases (Kroon et al. 1996; Ralet et al. 1994; Williamson et al. 1998a). The type B feruloyl esterases show a preference for the phenolic moiety of the substrate that contains one or two hydroxyl substitutions, as found in p-coumaric and caffeic acids, respectively. Hydrolytic rates are significantly lowered when a methoxy group is present. Regarding specificity against synthetic substrates, type B feruloyl esterases are active against methyl caffeate, methyl ferulate and methyl p-coumarate, but not methyl sinapate.

A third type of esterase

A third type of esterase with broad specificity against synthetic hydroxycinnamic acids (ferulic, caffeic, p-coumaric, sinapic) has been reported as an acetyl esterase (XYLD) from Pseudomonas fluorescens subsp. cellulosa (Ferreira et al. 1993). The data indicate that XYLD is able to cleave ester linkages in plant cell walls, although the enzyme exhibits greatest activity towards the release of acetic acid from 4-nitrophenyl acetate. The hydrolysis rate of XYLD esterase, both for hydroxycinnamic acids and for acetylated substrates, is significantly lower (25- to 500-fold) than reported for the characterised fungal feruloyl esterases (De Vries et al. 2002b; Faulds and Williamson 1994; Kroon et al. 1996, 2000) and other microbial acetyl xylan esterases (Basaran and Hang 2000; Blum et al. 1999; Degrassi et al. 2000; Linden et al. 1994; McDermid et al. 1990). In addition, XYLD is capable of releasing 5-5′-ferulic dehydrodimers from barley and wheat cell walls (Bartolome et al. 1997). In respect of this broad substrate specificity, XYLD has been tentatively classified as a general plant cell wall esterase.

Feruloyl esterases are subject to complex regulation

The expression of many fungal genes coding for cell wall-degrading enzymes are subject to complex regulation through carbon catabolite repression, as exercised by the repressor CreA from A. niger and A. nidulans (De Vries et al. 1999, 2002a; Dowzer and Kelly 1991; Orejas et al. 2001; Ruijter and Visser 1997), Cre-1 from N. crassa (Ebbole 1998), or Cre1 from Trichoderma reesei (Strauss et al. 1995). Cell wall-degrading enzymes of A. niger are also specifically induced via the xylanolytic transcriptional activator XlnR (De Vries and Visser 1999; van Peij et al. 1998), in addition to positive or negative regulation by factors derived from the growth substrates themselves (De Vries et al. 2002b; Faulds and Williamson 1999; Faulds et al. 1997). Feruloyl esterase activity is proposed to be regulated through the xylose-induced expression of the repressor protein XlnR in Aspergillus during growth on hemicellulosic material (De Vries and Visser 2001), but it has yet to be established how type B esterases are induced during growth on material such as sugar beet pulp.

Protein sequence databases

In recent years, the number of microbial feruloyl esterase activities reported has increased, in particular with the acquisition of related protein sequences in the growing genome databases. Several enzymes have been purified and characterised from aerobic and anaerobic micro-organisms that utilise plant cell wall carbohydrates (Blum et al. 2000; Borneman et al. 1992; Castanares et al. 1992; Crepin et al. 2003a; Dalrymple et al. 1996; De Vries et al. 2002b; Donaghy et al. 2000; Faulds and Williamson 1991, 1994; Ferreira et al. 1993; Fillingham et al. 1999; Kroon et al. 1996, 2000; McCrae et al. 1994; Tenkanen et al. 1991). Feruloyl esterase activities have also been detected in mammalian cells and plants (Andreasen et al. 2001; Sancho et al. 1999) but these enzymes have yet to be purified and their protein sequences determined for comparison with the microbial enzymes. However, a number of microbial gene sequences have been determined; and the predicted protein sequences from these suggests there is a considerable degree of structural diversity between them. Analysis of predicted feruloyl esterase protein sequences show that many of these enzymes are modular, comprising of a catalytic domain covalently fused to non-catalytic carbohydrate-binding modules (Ferreira et al. 1993; Kroon et al. 2000). Alternatively, the catalytic domain may be included in a greater modular complex, such as that observed for the cellulosome of Clostridium sp. (Blum et al. 2000; Shoham et al. 1999) or for the multiprotein cellulose-binding cellulase-hemicellulase complex from the anaerobic fungus Piromyces equi (Fillingham et al. 1999). Although the first crystal structure of a feruloyl esterase module was reported recently for two esterases from C. thermocellum (Prates et al. 2001; Schubot et al. 2001), relatively few studies have been performed to elucidate the functional relationships between sequence-diverse feruloyl esterases. At present, the lack of highly conserved sequences within the sequenced esterases does not permit further classification for the feruloyl esterases, other than that their primary amino acid sequences place them in family 1 (Coutinho and Henrissat 1999) of the carbohydrate esterase classification (http://afmb.cnrs-mrs.fr/~cazy/CAZY/).

Based on the primary sequence identity and activity profile data reported for more than 20 feruloyl esterases against synthetic methyl esters, we propose a scheme for the sub-classification of the feruloyl esterase enzymes. The classification is discussed in terms of the evolutionary relationships existing between carbohydrate esterases.

Materials and methods

Phylogenetic analyses

Multiple alignments of sequences or domains showing feruloyl esterase activity and sequence-related enzymes, such as lipases, and the construction of a neighbour-joining phylogenetic tree (Saitou and Nei 1987) were performed with the CLUSTAL W program, available at http://www.ebi.ac.uk/clustalw/ (Thompson et al. 1994). The matrix used for the multiple sequence alignment was the group of BLOSUM matrices recommended for amino acid sequences alignments (Henikoff and Henikoff 1992). The gap penalties were set at the default values, the gap separation penalty value was 8, the penalty value for extending a gap was 0.05 and the penalty value for opening a gap was 10.

Results and discussion

Feruloyl esterase sub-classification

Using literature reports of the substrate preferences of characterised feruloyl esterases, together with the growth substrate requirements for crude cell wall materials leading to the expression of these enzymes by various microbes, we organised the microbial feruloyl esterases into four functional classes. Using the published compatible data on the feruloyl esterases Pen. funiculosum FaeB (EMBL accession number AJ291496), N. crassa Fae-1 (AJ293029), Ps. fluorescens XYLD (X58956), Pen. funiculosum FaeA (AJ312296), Piromyces equi EstA (AF164516), A. niger FaeA (AF361950); Talaromyces stipitatus FaeC (AJ505939) and A. niger FaeB (AJ309807), a comparative analysis indicated these enzymes could be divided into four putative sub-classes, termed types A, B, C and D (Table 1). Members of each sub-class show similar activity profiles against four model substrates [methyl 3-methoxy-4-hydroxycinnamate (MFA), methyl 3,4-dihydroxycinnamate (MCA), methyl 3,5-dimethoxy-4-hydroxycinnamate (MSA), methyl 4-hydroxycinnamate (MpCA)] and similar abilities to release diferulate or not from plant cell wall material. Moreover, these classes are likely to reflect the substrate availability of natural plant cell walls, as they are preferentially induced by their respective organisms when grown in the presence of similar substrates. Finally, it can be demonstrated that members of the same activity class also share a high degree of amino acid sequence identity.

Table 1 Classification of feruloyl esterases based on primary amino acid sequence identity, specificity for hydroxycinnamic acid methyl esters, ability to release 5,5′-diferulic acid from model and complex substrates and inducible plant cell wall materials. CE carbohydrate esterase; for other abbreviations, see Materials and methods
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In order to investigate further the feruloyl esterase sub-classification, a phylogenetic tree was drawn on the basis of primary sequence identity between selected plant cell wall-acting esterases and other sequence related enzymes, such as lipases and xylanases. A phylogenetic tree of the cladogram type was constructed by the neighbour-joining method, using CLUSTAL W. A cladogram is a branching tree assumed to be an estimate of a phylogeny where the branches are of equal length, which shows common ancestry but does not indicate the degree of evolutionary time separating the taxa. The phylogenic tree indicates that feruloyl esterases diverged in three main branches, within which they evolved along discrete lines to form sub-classes of specialised enzymes (Fig. 1).

Fig. 1
figure 1

Cladogram of enzymes of the feruloyl esterase family and relatives. Sequence names with their accession numbers are shown on the right of the tree. NcFaeD-3.660 Neurospora crassa FaeD-3.660 feruloyl esterase, PfFaeA Penicillium funiculosum FaeA (AJ312296), PfXYLD Pseudomonas fluorescens XYLD (X58956), NcFaeD-3.544 N. crassa FaeD-3.544 feruloyl esterase (this work), PeESTA Piromyces equi ESTA (AF164516), NcFae-1 N. crassa Fae-1 (AJ293029), PfFaeB Pen. funiculosum FaeB (AJ291496), AaAXE Aspergillus awamori AXE (D87681), AnAXE Asp. niger AXE (A22880), AfAXE Asp. ficuum AXE (AF331757), PpAXEI Pen. purpurogenum AXE I (AAM93261), OspFaeA Orpinomyces sp. PC-2 FaeA (AF164351), CtXynZ Clostridium thermocellum XynZ (M22624), RspXyn1 Ruminococcus sp. Xyn1 (S58235), RfXynE R. flavefaciens XynE (AJ272430), CtXynY C. thermocellum XynY (X83269), RaXynB R. albus XynB (AB057588), RfAXE R. flavefaciens AXE (AJ238716), OspAXE Orpinomyces sp. PC-2 AXE A (AAC14690), AnFaeB Asp. niger FaeB (AJ309807), TsFaeC Talaromyces stipitatus FaeC (AJ505939), AoEST Asp. oryzae selective esterase (CAD28402), AspChloro Acinetobacter sp. chlorogenate esterase (AAL54855), AoTAN Asp. oryzae tannase (JC5087), RfXynB R. flavefaciens XynB (Z35226), AnFaeA Asp. niger FaeA (AF361950), AaFaeA Asp. awamori FaeA (Q9P979), AtFaeA Asp. turbingensis FaeA (Y09331), TlLipase Thermomyces lanuginosus lipase (O59952), RmLipase Rhizomucor miehei lipase (P19515), PpAXE P. purporogenum xylan esterase II (AF015285), TrAXE T. reesei acetyl xylan esterase I (S71334), BfCINI Butyrivibrio fibrisolvens E14 cinnamoyl ester hydrolase I (T44624), BfCINII B. fibrisolvens E14 cinnamoyl ester hydrolase II (AAB57776), NpAXE Neocallimastix patriciarum AXE (U66253), ApESTA Asp. parasiticus ESTA esterase (AF417002), AspAEST Acinetobacter sp. AEST esterase (BAB68337), BpAXE Bacillus pumilus AXE (AJ249957), SlAXE Streptomyces lividans AXE (AAC06115)

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The phylogenetic analysis by and large supports the feruloyl esterase classification proposed above on the basis of enzyme activity profiles against synthetic methyl esters. However, it is of note that the phylogenetic analysis indicates that TsFaeC (EMBL accession number AJ505939) and AnFaeB (AJ309807) are closely related and evolved from a common root. As a consequence, we suggest that these enzymes belong to the same sub-class of feruloyl esterases, termed type C, despite the reported difference in the substrate specificity of AnFaeB for methyl hydroxycinnamates, which initially suggested this enzyme to be a type B feruloyl esterase (De Vries et al. 2002b). The true type B enzymes have a common node, but the protein sequences of Pen. funiculosum FaeB (AJ291496) and N. crassa Fae-1 (AJ293029) may be distinguished from the high degree of conservation observed between the acetyl xylan esterases of A. awamori (D87681), A. niger (A22880), A. ficuum (AF331757) and Pen. purpurogenum (AAM93261).

The phylogenetic tree also recognises sequence relationships outside the present scheme. For example, the acetyl xylan esterase II from Pen. purpurogenum (Accession number AF015285) and the catalytic domain of AXE I (acetyl xylan esterase) from Trichoderma reesei (S71334) share 67% sequence identity (Gutierrez et al. 1998); and enzymes such as XynY and XynZ from C. thermocellum (Accession numbers P51584 and M22624, respectively), FaeA from Orpinomyces sp. PC-2 (AF164351) and Xyn1 from Ruminococcus sp. (S58235) are closely related (Blum et al. 2000). In parallel, the phylogenetic tree shows that, although the feruloyl esterases have some common roots, in some cases they may be more sequence-related to a variety of other enzymes, such as lipases, acetyl xylan esterases, or xylanases, than they are to each other.

Although the four enzyme sub-classes could be ascribed based on functional criteria outlined above, the phylogenetic analysis also suggests that there are probably yet further unnamed enzyme sub-classes. This fifth sub-class is comprised of OspFaeA (AF164351), CtXynZ (M22624), RspXyn1 (S58235), RfXynE (AJ272430), CtXynY (P51584) and RaXynB (AB057588). However, at present, this grouping may only be discriminated on the basis of their primary amino acid sequence identity, as no further correlation could be established here due to the lack of comparable enzyme activity data and, in some cases, information as to which plant cell wall materials preferentially provoke the expression of these feruloyl esterases by the microbes that possess them. Future acquisition of protein and predicted protein sequences with complementary enzyme activity data using synthetic methyl esters will enable the substantiation of these and potentially other enzymes classes recognised in phylogenetic analyses. These data will in turn provide a better understanding of the complex evolutionary relationship between the feruloyl esterases.

In conclusion, we propose that feruloyl esterases be classified into four functional sub-classes, termed types A, B, C and D, based on similarities in esterase activity profiles against synthetic methyl esters and underpinned by the intra- and inter-group protein sequence identities of the enzymes.