Identification and Characterization of Feruloyl Esterases Produced by Probiotic Bacteria

A variety of phenolic compounds are naturally available, and contain one or more phenolic rings with or without substituents such as hydroxyl or methoxy groups. The term phytophenol, or phytochemical, is also used due to the widespread distribution of these chemicals throughout the plant kingdom (Huang et al., 2007). Phytophenols are secondary metabolites of plants, which are primarily used in defense against ultraviolet radiation and pathogens (Beckman, 2000). These chemicals also participate in the formation of macromolecular structures in plant cell walls, and are naturally present in the form of monophenols or polyphenols with ester linkages. The presence of phenolic ester linkages limits the hydrolytic activity of enzymes such as xylanases, cellulases, and pectinases, by shielding the site of hydrolysis on plant cell walls from these enzymes. Hydrolyzing the ester linkages within the phytophenols releases the phenolic acids and relaxes the structure of the plant cell wall, aiding in the degradation and maximizing the nutritional value of dietary fiber. Phenolic acids such as ferulic acid, caffeic acid, chlorogenic acid, and rosmarinic acid are studied extensively due to their anti-oxidative, anti-inflammatory, and other health related properties which have been demonstrated both in vitro and in vivo (Srinivasan et al., 2007). Even though phenolic acids can be easily found in dietary fiber, the ester linkages prevent their absorption in the human intestine. It has been demonstrated that only small monophenolic acids, but not esterified phenolic acids, can be absorbed efficiently by the monocarboxylic acid transporter (Konishi et al., 2005). Thus, an enzymatic step is required to convert the esterified phenolic acids into monophenolic acids prior to absorption. In the presence of water, a specific type of enzyme, feruloyl esterases (FAEs), is able to hydrolyze the phenolic compounds into respective alcohols and phenolic acids. Thus, FAEs become one of the target fields of study to improve the bioavailiability and assimilation of phenolic acids in the human diet.

phenolic/aromatic form, such that an aromatic hydrocarbon is attached to the carbon atom of the carbonyl group of the ester. FAEs are important enzymes in the rumen ecosystem due to their ability to increase the absorption of plant-based energy sources in ruminant animals. In recent years, several FAEs from fungi were partially characterized, but little is known about bacterial or plant FAEs. For both medicinal and industrial applications, there is an increasing amount of research focused on FAEs that are capable of releasing monophenols from plant biomass. To the best of today's knowledge, humans do not synthesize FAEs. However, FAE activity is found in total human gut microbiota (Kroon et al., 1997, Gonthier et al., 2006, indicating that FAEs are present in the intestine and may contribute to the release of phytophenols from dietary fiber. Currently, most characterized FAEs have been identified from fungi, and the lack of FAEs identified in other organisms, particularly intestinal bacteria, has limited their application.
Tertiary structure information on FAEs is scarce, while primary and secondary structure is poorly conserved between fungi and bacteria. Identifying and characterizing FAEs in bacteria is an important challenge. This chapter tells the story of the identification, purification, characterization, and crystallization of FAEs from probiotic bacteria. The potential FAEs were identified based on the enzymatic activity displayed by the bacterial strains as well as bioinformatics analysis. The work discussed herein will provide insight for further exploration of FAEs in other species, enhancing the path for medicinal and industrial application of these enzymes.

Identification of feruloyl esterases from bacteria
Our understanding of the relevance of the commensal microbiota in relation to the healthy status of the host is rapidly expanding. However, the mechanisms by which these microorganisms interact with the host are still unclear. Important technological advances such as rapid sequencing methods, bioinformatics, and species identification using 16S rDNA are valuable tools to describe the variability and composition of these small ecosystems. One of the most interesting applications of the study of commensal microbiota is the identification of species potentially responsible for, or correlated with, specific host diseases. For example, there are noticeable changes in the composition of the gut microbial ecosystem of diabetes patients compared to healthy individuals (Vaarala et al., 2008). Some studies indicate that there is a predominant presence of probiotic bacteria such as Lactobacillus johnsonii, Lactobacillus reuteri, and Bifidobacterium species in healthy individuals (Roesch et al., 2009).
Lactic acid bacteria (Lactobacilli) are well known bacteria present in the human intestine and used in probiotic supplements. There are a variety of explanations in the literature as to the mechanisms responsible for the probiotic effects of these bacteria. These mechanisms include competitive exclusion of pathogens, secretion of bioactive compounds, immune system alteration, and host metabolism modification. However, there is no general consensus as to the mechanism of probiotics action, and studies of mechanism typically differ depending on the species or strain of bacteria. For example, a feeding study using the biobreeding diabetes-prone (BB-DP) rat model for type 1 diabetes with the intestinal bacteria L. johnsonii showed that oral administration of the probiotic bacterium L. johnsonii decreases the incidence of type 1 diabetes, possibly by decreasing the intestinal oxidative stress response (Valladares et al., 2010). The decreased oxidative stress at the intestinal level www.intechopen.com could be a consequence of multiple factors. For example, the rat chow is formulated with many ingredients containing 6% to 8% (weight to weight) of fiber in the form of sugar beet pulp. The sugar beet pulp is an important source of ferulic acid, a phytophenol with antioxidative and anti-inflammatory effects (Couteau et al., 1998). It has been demonstrated that low dosage of cinnamic acids (especially ferulic acid) has been related with the stimulation of insulin secretion (Balasubashini et al., 2003, Adisakwattana et al., 2008, prevention of oxidative stress and lipid peroxidation (Balasubashini et al., 2004, Srinivasan et al., 2007, and inhibition of diabetic nephropathy progression (Fujita et al., 2008). The interaction of select bacteria in the host intestines with dietary fiber, and the possible release of phenolic acids, is an interesting process to be analyzed in order to generate a rationale understanding of the problem.
Despite the fact that total human gut microbiota displays FAE activity, specific bacterial species producing FAEs have not been investigated in depth. FAE activity was identified in several lactobacilli, including L. fermentum, L. reuteri, L. leichmanni, and L. farciminis, however the genes encoding the FAEs were not identified (Donaghy et al., 1998). It is hypothesized that probiotic bacteria could enhance the release of bioactive phenolic acids from dietary fiber by producing the necessary FAE activity and aiding in the assimilation of phenolic acids. It is necessary to identify and characterize the FAEs from probiotic bacteria to further investigate this hypothesis.

Identification of FAE-producing bacterial strains
A screening assay for detection of FAE activity from Lactobacillus strains was first described by Donaghy et al. (Donaghy et al., 1998). A model substrate for FAEs, ethyl ferulate, was embedded in de Man Rogosa Sharpe (MRS) plates. The presence of ethyl ferulate created a turbid appearance in the MRS agar due to the semi-soluble ethyl ferulate at 0.1% (weight to volume) final concentration. Ferulate assay (MRS-EF) plates were inoculated with cells obtained from individual overnight MRS cultures. The plates were incubated at 37°C in a gas pack system for a maximum of 3 days. The formation of halo (clear area) around the colonies due to the hydrolysis of ethyl ferulate indicated the presence of FAE activity. This technique was sucessfully applied by Lai et al. (Lai et al., 2009), which identified that L. johnsonii, L. reutri, and L. helveticus are able to produce FAEs ( Fig. 1).

Selection of potential FAEs using bioinformatics analysis
Since FAEs are classified as carboxylic esterases, they display the characteristics of serine esterases (Brenner, 1988, Cygler et al., 1993. These enyzmes have a classically conserved pentapeptide esterase motif with a consensus sequence glycine-X-serine-X-glycine (GlyXSerXGly), where X represents any amino acid. Public databases such as Comprehensive Microbial Resource (CMR) Database (Davidsen et al., 2010) can be used to predict potential FAEs from the genomic sequences of FAE producing strains. Since there are only a handful of bacterial FAEs currently identified, most of the potential FAEs are annotated as hypothetical hydrolases or esterases. Two FAEs, LJ0536 and LJ1228 were previously identified in a probiotic bacterium L. johnsonii (Lai et al., 2009). The amino acid sequences of both FAEs can also be used to identify potential FAE homologs using a BLASTP search in NCBI database (Altschul et al., 1997).

Purification and characterization of FAEs
Apart from fungal FAEs, most of the bacterial FAEs described in the literature are purified directly from the growth media of bacterial strains, without previous knowledge of their coding gene or protein amino acid sequence. In order to obtain a large amount of enzyme to carry out thorough biochemical characterization, it is necessary to express and purify the target FAEs as recombinant proteins.

Cloning and purification of FAEs
After selection of potential FAEs from genomic analysis of FAE-producing strains, the genes of interest can be cloned into an expression vector. pET vectors are one of the most common expression vectors used for the cloning and expression of recombinant proteins in Escherichia coli. The pET System is driven by the T7 promoter, so target genes are regulated by the strong bacteriophage T7 transcription and translation signals. In this system, gene expression is effectively induced by the presence of isopropyl -D-1-thiogalactopyranoside (IPTG). The host cells, such as E. coli BL21 DE3, provide T7 RNA polymerase during protein expression. The proteins are expressed with specific tags such as Histidine Tag or S epitope tag, depending on the vector used.
Using pET15 vector as an example, the expression of His 6 -tagged proteins is carried out in E. coli BL21 using IPTG (1 mM) to induce gene transcription on the recombinant vector p15TV-L. The cells are collected by centrifugation at 8000 RPM (JLA8.1000 rotor, Beckman Coulter) for 25 min. The collected cell mass is resuspended in 25 mL binding buffer (5 mM imidazole, 500 mM NaCl, 20 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) pH 7.5) and then disrupted by French press (20000 psi). The cell free extract is collected by centrifugation at 4°C, 17500 RPM (JA25.50 rotor, Beckman Coulter) for 25 min. The soluble His 6 -tagged proteins are purified by affinity chromatography. All solutions pass through the nickel-nitriloacetic acid (Ni-NTA) column by gravity flow. The Ni-NTA column is first washed with 30 mL of ultra-pure water to wash out unbound nickel ions. It is then preequilibrated with 30 mL binding buffer. The cell free extract is then applied to the Ni-NTA column. During this step, the His 6 -tagged proteins bind to nickel ions that are immobilized by NTA. The resin is washed with 30 mL of binding buffer to wash out any unbound proteins. 200 mL of wash buffer (20 mM imidazole, 500 mM NaCl, 20 mM HEPES pH 7.5), which contains a higher concentration of imidazole, is used to remove unspecific proteins that are bound to the resin. Imidazole is a competitive molecule that displaces the nickel ions bound to His 6 -tagged protein. The His 6 -tagged proteins are eluted using 20 mL elution buffer (250 mM imidazole, 500 mM NaCl, 20 mM HEPES pH 7.5). The purified proteins are dialyzed at 4°C for 16 hours. The dialysis buffer is composed of 50 mM HEPES buffer pH 7.5, 500 mM sodium chloride (NaCl), and 1 mM dithiothreitol (DTT). After dialysis, the samples are flash frozen and preserved at -80°C in 200 μL aliquots until needed. The His 6tag can be removed by treatment with tobacco etch virus (TEV) protease (60 ug TEV protease per 1 mg of target protein) at 4°C for 16 hours. The sample is applied to the nickel affinity chromatography column to eliminate the released His 6 -tag. Collected proteins are dialyzed at 4°C against dialysis buffer for 16 hours. The purified proteins without His 6 -tag are flash-frozen and preserved in small aliquots at -80°C until needed.
A rapid method to evaluate the FAE activity can be used immediately after purification. 3 µl of the purified proteins (3-5 µl equivalent to 0.1 µg total protein) are dropped on the surface of the MRS-EF screening plate. The formation of halo zones indicates the presence of FAE activity. This system was used to purify the recombinant proteins LJ0536 and LJ1228 (Lai et al., 2009). By using the same strategy, a hypothetical protein LREU1684 was purified from L. reuteri and identified as a FAE.

Enzymatic substrate profile analysis
The change in pH that occurs during ester hydrolysis can be used to screen for substrate preference of FAEs. A pH indicator such as 4-nitrophenol (Janes et al., 1998) can be used to detect the change of pH during a reaction by monitoring the absorbance with a spectrophotometer. From the information in absorbance change, an estimate of enzymatic activity on different ester substrates can be determined. Enzymatic substrate profiles are determined at 25°C using an ester library composed of a variety of aliphatic and phenolic ester substrates (Liu et al., 2001). The purified enzymes are first thawed from -80°C and redialyzed against 5 mM N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid (BES) buffer pH 7.2. The reactions are carried out in 96 well plates. Each enzymatic reaction contains 1 mM ester substrate, 0.44 mM 4-nitrophenol (proton acceptor), 4.39 mM BES pH 7.2, 7.1% (volume to volume) acetonitrile, and 30 -35 µg . mL -1 of enzyme in a total volume of 105 μL reaction mixture. The 96 well plates are incubated at 25°C using Synergy™ HT Multi-Detection Microplate Reader (Biotek). The reactions are continuously monitored for 30 min at 404 nm. FAEs such as LJ0536 and LJ1228 display high activity towards phenolic esters (Lai et al., 2009).
This techique is used only to demonstrate enzyme substrate preferences since it allows the use of several substrates in parallel. It utilizes specific conditions to detect the release of hydrogen ion (proton) during hydrolysis. The buffer (BES buffer) and the pH indicator (4nitrophenol) to be used in this type of assay must have similar affinity (BES buffer pK a = 7.09; 4-nitrophenol pK a = 7.15) for the release of protons. In this way, the ratio of protonated buffer and the protonated indicator remains constant. The pH shifts by the proton release during the enzymatic reaction and it is detected as a change in the yellow color of the indicator present in the mixture. Thus, this technique is not flexible enough to adjust to the best conditions (pH, type of buffers, ions, etc.) that many enzymes require to function at their maximum activity. The specific enzyme activity determined using this www.intechopen.com method does not reflect the true specific enzyme activity. In addition, the stability of several enzymes could be affected by exhaustive dialysis in the reaction buffer. The dialysis was performed using 120 to 150 times the volume of the original enzyme suspension. Consequently, the technique is valid only to demonstrate substrate preferences. An alternate method involving the commerical equipment HydroPlate ® instead of the tranditional 4-nitrophenol pH indicator can be used to monitor the pH during ester hydrolysis. The HydroPlate ® is a 96 well plate containing immobilized pH sensor layers in each well (PreSens). The sensor contains a stable reference dye and one sensitive to oxygen. Thus, the measured fluorescence used to determine pH are internally referenced for precision across the plate.

Biochemical properties of FAEs
As enzyme reactions are saturable, the biochemical parameters such as K m (Michaelis constant: amount of substrate required to reach half of V max ), V max (maximum rate of reaction or maximum enzyme specific activity, μmol . mg -1 . min -1 ), K cat (catalytic rate constant, s -1 ), and K cat / K m (catalytic efficiency, M -1 . s -1 ) can be determined by measuring the initial rate of the reaction over a range of substrate concentrations. The optimal conditions such as pH and temperature should be determined with model substrates before attempting to determine these biochemical parameters. Naphthyl esters and 4-nitrophenyl esters are common model substrates used to determine the biochemical parameters of esterases using saturation kinetics. Hydrolysis of naphthyl esters or 4-nitrophenyl esters generates napthol or 4-nitrophenol respectively, resulting in a specific absorbance at 412 nm. In similar ferulic acid esterase screenings, phenolic esters such as ethyl ferulate and chlorogenic acid should be included to compare the differences in biochemical parameters between aliphatic and phenolic esters. The hydrolysis of these phenolic esters can be monitored at 324 nm (Lai et al., 2009). FAEs have higher catalytic efficiency and affinity towards phenolic esters. However, phenolic esters have higher values of absorbance and are less stable compared to the model substrates such as 4-nitrophenyl esters. The concentration of phenolic esters used to obtain the initial rate of the reaction is usually limited to 0.1 mM due to the upper limit of plate reader absorbance. Using absorbance to obtain enzyme kinetic parameters from phenolic ester hydrolysis can be difficult. An alternative method such as high performance liquid chromatography could be used to estimate the enzymatic activity by monitoring the release of products during phenolic ester hydrolysis (Mastihuba et al., 2002).

Structural analysis of FAEs
FAEs belong to a structural group described as / fold hydrolases (Ollis et al., 1992). The secondary structure of this group is composed of a minimum of eight -strands in the center core surrounded by -helices. The term / barrel is also used to describe the structure. The -strands in the central core and -helices are mostly parallel. The -helices andstrands tend to alternate along the chain of the polypeptide. There are only few structures of FAEs deposited in public databases (PDB) (Berman et al., 2000). All the structures (apoenzymes or co-crystallized with a substrate) deposited in the PDB belong to two enzymes purified from only two species, a fungus Aspergillus niger and a bacterium Butyrivibrio www.intechopen.com proteoclasticus. Five additional structures related to the probiotic bacterium L. johnsonii FAE LJ0536 were recently released in the PDB. The recently available structures of bacterial FAEs will allow researchers to identify new FAEs based on structural alignments and conserved structural features.

GlyXSerXGly is the classical esterase motif
In general, carboxylic acid esterases have one classical esterase motif composed of a consensus sequence GlyXSerXGly. Analysis of the LJ0536 amino acid sequence showed that LJ0536 has two GlyXSerXGly motifs, an exception to the general one motif rule for carboxylesterases. Mutiple sequence alignments indicated that LJ0536 and its homologs, including LREU1684, all have two GlyXSerXGly motifs (Fig. 2). The reason for the presence of two GlyXSerXGly motifs in enzymes has not been addressed clearly in the literature. A recent structural study on LJ0536 (Lai et al., 2011) shows that LJ0536 displays a typical / hydrolase fold, which is composed of twelve -strands and nine -helices. Only one GlyXSerXGly motif (Gly104-X-Ser106-X-Gly108) of LJ0536 is catalytically active, while the other (Gly66-X-Ser68-X-Gly70) maintains the folding of the protein by hydrogen bonds. The newly identified FAE LREU1684 shares 47% amino acid sequence identity with LJ0536.

Identification of critical amino acids involved in substrate hydrolysis
Ferulic acid esterase features such as the catalytic triad and the oxyanion hole are usually maintained by several amino acid residues that are highly conserved among homologs. Other critical amino acids involved in substrate recognition and binding are also conserved among closely related homologs, but not necessarily with less related homologs. A technique called alanine scanning, or site-directed mutagenesis, is helpful to determine the conserved amino acids critical for catalysis in proteins with unknown structure. The target amino acids selected for modification are replaced by alanine. Alanine is chosen because the inert alanine methyl functional group generally does not interact with other residues or alter the overall protein structure. To introduce the alanine mutation, 39-nucleotide long complementary primers containing the desired amino acid replacement are used to introduce individual mutations. The protein variants are then constructed by Polymerase Chain Reaction using Finnymes Phusion TM high fidelity DNA polymerase. This approach was used to identify the critical amino acids of LJ0536 (Lai et al., 2011).
The enzymatic activities of alanine variants are impared when the mutated amino acids are critical to function of the proteins. However, the results obtained from alanine scanning may not be useful in distinguishing the specific function of the amino acids, such as the involvement of amino acids in the formation of catalytic triad, oxyanion hole, tertiary structure of the protein, or substrate recognition and binding. The amino acids involved in substrate recognition and binding can be determined by measuring the enzymatic activity of the protein variants with different substrate types. For example, mutation of the amino acids that are only necessary for phenolic ester binding would not impair the enzymatic activity when aliphatic esters are used as substrates (Lai et al., 2011). Ultimately, the tertiary structure of the proteins are still necessary to conclude the findings from alanine scanning.

Catalytic triad of FAEs is composed of serine, histidine, and aspartic acid
Two basic steps are involved during ester hydrolysis: acylation and deacylation (Ding et al., 1994). During acylation, the hydroxyl oxygen of the catalytic serine carries out a nucleophilic attack on the carbonyl carbon of the ester substrate. After the attack, a general base (the histidine of the catalytic triad) deprotonates the catalytic serine and the first tetrahedral intermediate is formed. The hydrogen bonding of the third member of the triad, aspartic acid, plays a critical role in the stabilization of the protonated histidine. The oxyanion of the resulting tetrahedral intermediate is positioned towards the oxyanion hole. The oxyanion hole is created by hydrogen bonding between the substrate carbonyl oxygen anion and the backbone of two nitrogen atoms from other residues of the catalytic pocket. The general base, histidine, transfers the proton to the leaving group. The deprotonation of histidine leads to the protonation of an ester oxygen to release the first product (for example: methanol with methyl ferulate as substrate). As a consequence, the tetrahedral intermediate collapses and the characteristic acylenzyme intermediate is formed. Thus, the residual half of the substrate remains attached to the catalytic serine.
The second step of the reaction, deacylation, takes place in the presence of water. A molecule of water performs a nucleophilic attack on the carbonyl carbon of the remaining substrate in the acylenzyme intermediate. The general base (histidine) immediately deprotonates a molecule of water, leading to the formation of a second tetrahedral intermediate. The catalysis follows a similar pattern described for the acylation. The second tetrahedral intermediate is stabilized by the formation of the oxyanion hole. The proton of the general base moves to the nucleophilic serine. Consequently, the ester oxygen is protonated and the tetrahedral intermediate collapses. The protonation of ester oxygen releases the final product (for example: ferulic acid with methyl ferulate as substrate), and reconstitutes the native serine residue and the original state of the enzyme.
The catalytic center of esterases always consists of a triad composed of a nucleophile (serine or cysteine), a fully conserved histidine, and an acidic residue (aspartic acid). In order for the catalytic triad residues to carry out their roles during hydrolysis as described above, the histidine must be located next to the catalytic serine, while the aspartic acid must be located next to the histidine. The catalytic triad of LJ0536 is composed of serine, histidine, and www.intechopen.com aspartic acid (Ser106, His225, Asp197). Due to the high amino acid sequence identity between LJ0536 and LREU1684, it is expected that both enzymes could have similar tertiary structures. Hypothetical tertiary structure of LREU1684 is predicted using SWISS-MODEL (Arnold et al., 2006). SWISS MODEL is a structure homology-modeling server, which allows users to predict the structure of a protein with a simple input of the peptide sequence. The modeling is generated based on the existing protein structures. The results indicate that the folding of LREU1684 is highly similar to LJ0536 (PDB: 3PF8). The catalytic triad of LREU1684 is arranged in an identical orientation as in LJ0536. It is composed of Ser109, His228, and Asp200 (Fig. 3). The catalytic serine residue (Ser109) is located on top of the sharp turn of an -helix (nucleophilic elbow). The catalytic triad arrangement of both LJ0536 and LREU1684 follows the general rule of ester hydrolysis.

Classical oxyanion hole aids in substrate binding
Co-crystallization assays of the LJ0536 catalytic serine deficient mutant Ser106Ala (LJ0536-S106A) with various ligands identifies the classical oxyanion hole of LJ0536 (Lai et al., 2011). LJ0536-S106A was co-crystallized with ethyl ferulate (PDB: 3QM1), ferulic acid (PDB: 3PFC), and caffeic acid (PDB: 3S2Z). All these structures show that the oxyanion hole of LJ0536 is formed by the backbone nitrogen atoms of phenylalanine and glutamine (Phe34 and Gln107). The oxyanion hole is an important structural feature, which stabilizes the tetrahedral intermediates during hydrolysis. Structural superimposition of LREU1684 and LJ0536 shows that the oxyanion hole of LREU1684 is formed by the backbone nitrogen atoms of Phe34 and Gln110 (Fig. 4).

Specific inserted domain contributes to substrate binding
The study of the LJ0536 structure indicated that a specific / inserted domain is critical for substrate binding (Lai et al., 2011). The inserted domain of LJ0536 is formed by a sequence of 54 amino acids from proline to glutamine (Pro131 to Qln184), and is located on top of the binding cavity. The two protruding hairpins from the inserted domain decorate the entrance and form the roof of the catalytic compartment. The phenolic ring of the ester substrate binds in the deepest part of the binding cavity, towards the inserted domain. In addition, three amino acid residues of the inserted domain, Asp138, Tyr169, and Gln145, contribute to the specific phenolic ester binding. The 4-hydroxyl group (ethyl ferulate, ferulic acid, and caffeic acid) and 3-hydroxyl group (caffeic acid) of the phenolic ring of the substrates are hydrogen bonded to Asp138 and Tyr169, respectively. Gln145 creates a bridge-like structure on top of the binding cavity, serving as a physical clamp holding the substrate inside the binding cavity. It also orients a water molecule in the binding cavity, which is important for activating the catalytic serine residue during hydrolysis.
Similar to LJ0536, LREU1684 has an / inserted domain formed by a sequence of 53 amino acids from Pro134 to Qln185 (Fig. 5). Asp141, Gln148, and Tyr172 of LREU1684 correspond to Asp138, Gln145, and Tyr169 of LJ0536, respectively. They adopted the same orientations as the residues in LJ0536 (Fig. 6). Thus, it is highly possible that Asp141, Gln148, and Tyr172 of LREU1684 also adopt the functional roles of Asp138, Gln145, and Tyr169 of LJ0536. LREU1684 and LJ0536 have both high amino acid sequence identity and high structural conservation.

Folding of LJ0536 is conserved among homologs
Since LREU1684 is predicted to have tertiary structure and binding mechanism that are similar to LJ0536, it is hypothesized that the other LJ0536 homologs should also contain the structural features of LJ0536. To test this hypothesis, the models of LJ0536 homologs are predicted using SWISS MODEL. The quality of the modeling is estimated by the E-value, QMEAN Z-Score, and QMEANscore4 (Benkert et al., 2011). The E-value is a parameter that describes the number of hits that you expect to find a protein by chance when searching a database. The lower the E-value, the more structurally significant the hit is. The Q-MEAN Z-Score measures the absolute quality of a model. A strongly negative value indicates a model of low quality. The QMEANscore4 represents the probability that the input protein matches the predicted model. The value ranges between 0 and 1. The results obtained using an automatic template search are summarized in Table 1.
All predictions provided good quality models except for the modeling of EVE, a hypothetical protein from Eubacterium ventriosum ATCC 27560. EVE has an E-Value of 1.40E-28, a QMEANscore4 of 0.477, and a QMEAN Z-Score of -4.276. BFI-1, a cinnamoyl ester hydrolase from Butyrivibrio fibrisolvens E14, has the best quality of model with an E-Value of 1.61E-91, a QMEANscore4 of 0.82, and a QMEAN Z-Score of 0.425. Among all 12 homologs, 10 were predicted to have similar folding to Est1E (Goldstone et al., 2010), a feruloyl esterase from Butyrivibrio proteoclasticus (PDB: 2wtmC and 2wtnA). The structures of LJ0536 and Est1E are highly similar as previously studied (Lai et al., 2011). The predictions were validated by including the sequences of LJ0536 in the analysis. The homologs, LBA-1 and BFI-2, do not have a similar Est1E folding. LBA-1 is annotated as / superfamily hydrolase in L. acidophilus NCFM. It was predicted to be similar to lipase www.intechopen.com in Burkholderia cepacia (PDB: 1YS1). BFI-2 is annotated as a cinnamoyl ester hydrolase in B. fibrisolvens E14. It was predicted to be similar to acetyl xylan esterase in Bacillus pumilus (PDB: 3FVR). In order to prove that the folding of LJ0536 is conserved in LBA-1 and BFI-2, a second prediction was preformed using Est1E or LJ0536 as the template structure. When Est1E was used as the template, the E-value of LBA-1 improved from 2.40E-08 to 2.70E-32. QMEAN Z-Score and QMEANscre4 decreased from -2.414 to -3.495 and from 0.556 to 0.527, respectively. When the prediction was done using LJ0536 as a template, the E-value improved to 1.2E-32, the QMEAN Z-Score decreased to -2.533, and the QMEANscre4 improved to 0.598. A similar scenario was observed when the protein BFI-2 was analyzed.

Protein
The results indicated that the folding of LJ0536 is conserved in LBA-1 and BFI-2. The overall structure of LJ0536 is conserved among all homologs studied. www.intechopen.com

Conclusion
FAE application is one of the major fields of study for improving the bioavailability of phenolic acids in food components (phytophenols). After FAE activity on phytophenols in the intestinal tract, released phenolic acids become bioavailable and are absorbed in the intestines and can provide beneficial effects to the host. The identification and crystallization of the first intestinal probiotic bacterium FAE, LJ0536 identified from L. johnsonii, provides the fundamental knowledge (protein sequence and structural features) required to further identify FAEs from other species. Using a hypothetical protein LREU1684 as an example, this chapter provides a basic approach on how to identify, purify, and characterize FAEs, predict the model structure, and compare the model with known FAE structures. Further FAE crystallization is required to prove that the structure of LJ0536 is conserved among all homologs.