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Plant Cell Tiss Organ Cult (2017) 130:13–24 DOI 10.1007/s11240-017-1200-0

ORIGINAL ARTICLE

Horseradish esterases: detection, purification and identification

Ivana Leščić Ašler1 · Petra Peharec Štefanić2 · Biljana Balen2 · Günter Allmaier3 · Martina Marchetti‑Deschmann3 · Biserka Kojić‑Prodić1 

Received: 17 August 2016 / Accepted: 10 March 2017 / Published online: 19 March 2017 © Springer Science+Business Media Dordrecht 2017

Keywords Armoracia lapathifolia · Protein identification · Protein purification · Electrophoresis · Mass spectrometry · Esterase

Introduction

In vitro grown tissues can be used for production of biolog-ically active compounds. Esterases are known to participate in many biological processes in plants, but only a handful of plant esterases are thoroughly characterized. For many decades now, hydrolases (esterases in particular) have been studied as valuable tools for stereospecific hydrolysis and synthesis of many biotechnologically important com-pounds. Microbial enzymes have gained interest for their wide uses in bioconversions related to food, agriculture, production of fine chemicals, medicine, diagnostics, energy and waste treatment (Singh et al. 2016; Gurung et al. 2013; Sayali and Surekha 2013). Esterases have been used in synthesis of chiral drugs, in producing flavoring and fra-grance compounds, as well as for hydrolysis of milk fat for the purpose of flavor enhancement in the manufacturing of cheese-related products (Panda and Gowrishankar 2005). Microbial esterases from baker’s yeast and aroma esters have been used in production of alcoholic beverages, over three decades (Suomalainen 1981). The research related to enzymatic degradation of acetyl-xylan by cooperativity of fungal esterases and xylanases, published in 1985 (Bieley et  al. 1985), initiated very important studies on decom-position of biomass which is among the highest priority in biotechnology these days (Liu and Ding 2016). Bacte-rial and fungal esterases are generally well-characterized and widely applied (Borelli and Trono 2015), due to the simple access to large amounts of these enzymes (since genetic manipulation with enzymes of microbial origin is

Abstract Our goal is to characterize esterases from horseradish tissues and assign their physiological roles. In the present study we focused on isolation, purification and identification of esterases from different horserad-ish tissues: plantlets and two tumor tissue lines. Horizon-tal IEF system enabled separation of six esterase isoforms with quite different pI values as well as with pronounced differences in expression levels among analyzed tissues. Esterases were extracted, fractionated by means of cation exchange chromatography, and analyzed by planar gel elec-trophoresis (SDS–PAGE) and isoelectrical focusing (IEF), UV/Vis spectroscopy, MALDI mass spectrometry (MS) and MALDI-MS/MS. Several chromatographic strategies were applied for esterase purification and characteriza-tion. Two subsequent cation exchange chromatographic steps based on SP-Sepharose FF material, followed by in-solution digestion combined with MALDI-MS and MS/MS proved to be the best strategy for identification of two ester-ase proteins, namely Pectinesterase/pectinesterase inhibitor 18 and GDSL esterase/lipase ESM1.

* Ivana Leščić Ašler ilescic@irb.hr

1 Department of Physical Chemistry, Laboratory for Chemical and Biological Crystallography, Ruđer Bošković Institute, Bijenička cesta 54, 10000 Zagreb, Croatia

2 Division of Molecular Biology, Department of Biology, Faculty of Science, University of Zagreb, Horvatovac 102a, 10000 Zagreb, Croatia

3 Institute for Chemical Technologies and Analytics, Technische Universität Wien (Vienna University of Technology), Getreidemarkt 9/164, 1060 Vienna, Austria

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straightforward and fast). However, plant esterases were not so much in focus of research during the last years. Only a few plant esterases are biochemically well-characterized and mere 20 structures of 11 plant esterases are deposited in the Protein DataBank (PDB, March 2016). To date, among the hydrolases from plant sources, only cabbage phospholipase D has been successfully applied to indus-trial processes (Borelli and Trono 2015). Still, it is known that esterases are involved in regulating many biological processes in plants (Gershater and Edwards 2007). Per-haps most widely studied in this respect are pectinmethyl esterases that participate in developmental processes (cell division, expansion and differentiation) by altering the cell wall properties through pectin demethylation (Bou Daher and Braybrook 2015). However, GDSL family of esterases/lipases were also found to have important roles in develop-ing seeds (Chen et al. 2012), during post-germination and seedling development in Arabidopsis (Huang et al. 2015); and during seed germination and morphogenesis in Bras-sica napus (Clauss et  al. 2008, 2011), while physiologic functions in coleoptile elongation and plant growth in the seedling stage were suggested for two rice GDSL esterases/lipases (Riemann et al. 2007; Park et al. 2010). Moreover, it has been reported that the carboxylesterase family partici-pates in olive pollen germination, pollen tube growth and penetration of the stigma (Rejón et al. 2012); while Schil-miller et  al. (2016) found that acylsugar acylhydrolases catalyze hydrolysis and remodeling of acylsugars in tomato trichomes, which have an important role in plant defense against herbivores.

Some plant esterases are known to regulate plant devel-opment and defense response through modification and activation of important signal molecules such as salicylic acid (Forouhar et al. 2005), jasmonic acid (Stuhlfelder et al. 2004) and auxin (Yang et al. 2008). Furthermore, esterases have roles in regulating plant bioactive metabolism (e.g. fruit ripening), bioactivation of herbicides, and detoxifi-cation as well as regulation of turnover of stored natural products (Jolie et  al. 2010). Even small changes in ester-ase activity represent a reliable indicator of metabolism changes (Vitacek et al. 2007).

ESTHER database (Lenfant et  al. 2013), compris-ing members of the alpha/beta-hydrolase superfamily together with the relatively new family of GDSL hydro-lases, contains 335 hydrolases from the model plant Arabidopsis thaliana (March 2016). The great number of family members, broad substrate specificity, stere-oselectivity and/or thermostability indicate great adapt-ability of organisms during evolution and the importance of this particular enzyme family for organism survival; which gives great possibilities for development of applied

biocatalysis (Grogan 2011). The recent gain of knowl-edge on eukaryotic enzymes (e.g. physico–chemical properties, optimal functional conditions, mechanism of action) and on means of their manipulation (e.g. highly successful eukaryotic expression systems, protein engi-neering methods) provides potential to expand possi-bilities of their application (Liu et  al. 2004; Borelli and Trono 2015). It is therefore of great importance to further study plant esterases in depth.

Plant tissue culture techniques have become one of the main tools in investigation of many unsolved problems in basic and applied sciences. They enable research on many levels (molecular, cell, organism) and can be used for production of biologically active compounds. One exam-ple is the culture of horseradish (Armoracia lapathifolia Gilib.), a common seasoning with medical applications (Nhat et al. 2013). Horseradish belongs to the same fam-ily as A. thaliana (Brassicaceae). However, its genome is not sequenced and differences among sequences of homologous proteins can be expected.

Horseradish in vitro culture comprises tissues of vari-ous differentiation levels and morphologies (Balen et al. 2009). Namely, in an attempt to obtain transformed horseradish plants, leaf explants of the plantlets were transformed with a wild strain B6S3 of Agrobacterium tumefaciens (Krsnik-Rasol and Muraja-Fras 1993). Sub-cultured on hormone-free MS medium (Murashige and Skoog 1962); the transformed tissues continued grow-ing as two different tumor lines: either as a completely unorganized tissue that never expressed any morphogenic capacity (TN—tumor tissue) or as teratogenous tumor (TM—teratoma tissue) with malformed hyperhydric shoots. All in  vitro-grown horseradish tissues already exhibited biotechnological potential (Krsnik-Rasol et  al. 1992; Redovniković Radojčić et  al. 2008) and therefore have been chosen as models for the research presented. Esterase activity has already been investigated using in  vitro grown plantlets, TM and TN by Krsnik-Rasol et  al. (1999), who demonstrated differences in expres-sion of certain esterase isoforms in leaves of horseradish plantlets compared to two tumor lines. Moreover, the iso-enzymes of esterase were shown to be useful biochemi-cal markers of developmental processes in sugar beet cell lines (Krsnik-Rasol et al. 1999), cactus Mammillaria gra-cilis in vitro-grown tissues (Balen et al. 2003), and Fritil-laria meleagris L. bulb-scale culture (Petrić et al. 2015).

Based on these results, we set out to identify and char-acterize esterases in horseradish tissues of different mor-phology levels to determine their physiological function. The developed purification strategy allowed as result iso-lation, identification and characterization of two isoforms of horseradish esterase.

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Materials and methods

Chemicals

Buffer chemicals were purchased from Kemika (Zagreb, Croatia). All organic solvents were of analytical grade (Kemika). Ultrapure water from Simplicity System (Milli-pore, Billerica, MA, USA) was used. If not specified, other chemicals used were purchased from Sigma (St. Louis, MO, USA).

Plant material

Horseradish plantlets, teratoma and tumor tissues have been propagated in  vitro on a solid MS nutrient medium (Murashige and Skoog 1962) containing 8.5 g L−1 of agar and 30  g  L−1 glucose without any growth regulator. Cul-ture conditions were: 24 °C, 16-h light/8-h dark period and irradiation of 33 μmol m−2 s−1. Tissues were subcultivated every 3 weeks. Fifteen days after being moved to the fresh nutrient medium, tissue explants were used for the extracts preparation. The transformed character of tumor and tera-toma cells had been confirmed by testing octopin produc-tion and PCR amplification of a 336 bp fragment of the 6a gene from TL-DNA (Peškan et  al. 1996). For subsequent analyses; horseradish plantlet leaves (L), teratoma (TM) and tumor (TN) tissues were used.

Soluble proteins were extracted from the leaves of 6–10 cm high plantlets and from TM and TN tissues in their exponential growth phase (9 days after subculturing). Tis-sue samples were homogenized in ice cold 100 mM potas-sium phosphate buffer, pH 7.0. Tissue mass (g) to buffer volume (mL) ratio was 1:3. To remove phenols, insoluble polyvinylpyrrolidone (cca 50 mg) was added to each tissue sample prior to grinding in the Omni Mixer Homogenizer 17150 (Omni International, NW Kennesaw, GA, USA) for 3 min at 30 Hz. The homogenates were centrifuged for 30 min at 20,500×g and 4 °C. Obtained supernatants were centrifuged again for 60 min at 20,500×g and 4 °C. Super-natants were collected and protein content was determined according to Bradford (1976) using bovine serum albumin (BSA) as standard.

Esterase activity assay

Esterase activity was measured towards 1- and 2-naphthyl acetate (1-NA and 2-NA, respectively; Serva, Heidelberg, Germany), broad spectrum carboxylesterase (EC 3.1.1.1) and arylesterase (EC 3.1.1.2) substrates. Stock solutions (0.1 M) of substrates were prepared in methanol. 1 mL of buffer (0.1  M Tris–HCl, pH 7.4) was mixed in a cuvette with 30 µL of methanol and 10 µL of substrate stock solu-tion. Absorbance at 322 or 313  nm (for 1-NA or 2-NA,

respectively) was measured and the reaction started by adding an appropriate amount of sample (5–100 µL). The increase in the absorbance was followed for 3 min. Ester-ase activity was corrected for spontaneous hydrolysis of substrates and expressed as µmol of produced alcohol per minute and mg of protein (µmol min−1 mg−1). For calcu-lating activities, the given molar absorption coefficients of 1-naphthol (Ɛ322nm = 2000  M−1  cm−1) and 2-naphthol (Ɛ313nm = 1250 M−1 cm−1) were applied (Balen et al. 2003).

For detection of esterases after horizontal isoelectric focusing (IEF), the gel was soaked for 30  min in a solu-tion comprised of 4  mg of 1-NA and 4  mg of 2-NA dis-solved in 1.6 mL of 50% acetone and mixed with 10 mL of 50 mM Tris–HCl buffer, pH 7.1. Afterwards, the gel was washed with dH2O and soaked for 20 min in 0.2% FastBlu-eRR salt solution (GE Healthcare, Piscataway, NJ, USA). Twenty mg of FastBlueRR salt was dissolved in 1 mL of methanol and mixed with 10 mL of 50 mM Tris–HCl buffer (pH 7.1) to obtain the above mentioned 0.2% solution. Fol-lowing this step, the gel was again washed with dH2O and fixed in 30% ethanol.

Protein concentration in extracts and combined chro-matographic fractions was routinely assayed according to Bradford (1976). When chromatographic fractions were tested for protein content, in order to save material and speed up the purification process, 10  µL of fraction was mixed with 100 µL of Bradford reagent in a ceramic 12-well plate and intensity of blue color estimated visually.

Chromatographic procedures

All chromatographic procedures were performed at 4 °C. Fractionation of proteins from horseradish tissues as the first purification step was performed by cation exchange chromatography on SP-Sepharose FF (GE Healthcare) column (53.3 mL). Extracts (typically 40 mL with protein concentration of 1.6  mg  mL−1) were first concentrated three times by ultrafiltration using an YM-10 membrane (Merck Millipore, Darmstadt, Germany) and transferred to 50 mM phosphate buffer (pH 7.0) by means of size-exclu-sion chromatography on Sephadex G-25 (GE Healthcare) column (113 mL). Cation exchange chromatographies were performed in 50  mM phosphate buffer (pH 7.0) and pro-teins eluted by stepwise increase in NaCl concentration (0, 0.2, 0.4, 0.6 and 1.0 M NaCl). Flow rate of 80 mL h−1 was maintained through chromatographic elution. With each NaCl concentration, 20 fractions of 5  mL were collected and tested for protein content. Fractions containing proteins were combined and concentrated by ultrafiltration through YM-10 membrane and tested again for protein content and esterase activity as previously described.

In the second purification step, part (3.5 mL with protein concentration of 0.4 mg mL−1) of fraction ‘b’ from cation

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exchange chromatography on SP-Sepharose FF (fractions eluted with 0.2 M NaCl) was transferred to 50 mM phos-phate buffer (pH 7.0) by means of the PD-10 column (GE Healthcare) and loaded onto smaller sized column of SP-Sepharose FF (7.3 mL). Chromatography was performed in the same buffer as the first one, with 60 mL h−1 flow rate, but proteins were eluted with 84 mL of NaCl gradient (0–0.3  M), followed with 25 mL of 1  M NaCl, and frac-tions of 0.84 mL were collected.

Chromatographic methods based on FPLC (fast protein liquid chromatography) system AKTA (GE Healthcare) were evaluated but gave no satisfactory result. One part (3.5 mL with protein concentration of 1.1 mg mL−1) of fraction ‘a’ from cation exchange chromatography on SP-Sepharose FF (unbound fractions, flow-through) was loaded onto anion exchange column MonoQ 5/50 GL and the second part (2.3 mL with protein concentration of 1.8 mg mL−1) of the same fraction was loaded onto size exclusion chro-matography column HiLoad 16/60 Superdex 200. MonoQ column was equilibrated in 50  mM Tris–HCl buffer (pH 8.0) and proteins were eluted with NaCl gradient (0–0.3 M in 30 mL, 0.3–1.0 M in 5 mL) and 1 M NaCl (5 mL) in the same buffer. Chromatography on Superdex 200 col-umn was performed in 20 mM Tris–HCl buffer with 0.2 M NaCl, pH 7.6. One part (3.5 mL with protein concentra-tion of 0.4 mg mL−1) of fraction ‘b’ from cation exchange chromatography on SP-Sepharose FF (fractions eluted with 0.2  M NaCl) was transferred to 50  mM phosphate buffer (pH 7.0) by means of the PD-10 column (GE Healthcare) and loaded onto cation exchange chromatography column MonoS 5/50 GL. This column was equilibrated in 50 mM phosphate buffer (pH 6.5) and proteins were eluted with NaCl gradient (0–0.3 M in 30 mL, 0.3–1.0 M in 5 mL) and 1  M NaCl (5 mL) in the same buffer. In all FPLC-based chromatographic separations the flow rate was maintained at 1 mL min−1 and 1 mL fractions were collected.

Electrophoretic methods

Horizontal isoelectric focusing (IEF) in one dimension was performed on PhastSystem by GE Healthcare, with pre-cast gels (pI 3–9) from the same producer, following the manu-facturer’s instructions. Protein pI markers (pI 3–10) were from the same company. Samples were loaded without any pretreatment. Gels were stained with esterase substrates 1-NA and 2-NA. Also, silver staining was used following the method of Hempelmann and Kaminsky (1986).

Horizontal sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS–PAGE) was also performed on Phast-System (GE Healthcare), with pre-cast gels (12.5%) and SDS buffer strips from the same producer, following the manufacturer’s instructions. Protein low molecular weight (LMW) markers (14.4–93.0  kDa) were from the same

company. Prior to loading, samples were mixed with treat-ment buffer (20 mM Tris–HCl (pH 8.0), 2 mM EDTA, 5% SDS, 10% 2-mercaptoethanol) in ratio 1:1 (v/v) and heated for 5 min at 95 °C in thermomixer (Thermomixer comfort, Eppendorf). Protein bands were visualized by silver stain-ing (Hempelmann and Kaminsky 1986).

Molecular weights of proteins in samples were deduced from linear regression based on electrophoretic migration of molecular weight markers, according to instructions from the manufacturer. Isoelectric points of proteins in samples were estimated using the pI markers, according to instructions from markers provider. Since pI markers were stained with silver and samples were stained with esterase substrates, both were not always loaded on the same gel. However, considering the reproducibility of the used IEF technique, estimated pIs were taken with confidence.

In‑solution digestion

Sample aliquots were dried, dissolved in 75 µL of denatur-ing buffer (6 M guanidine hydrochloride, 0.5 M Tris–HCl, 2 mM EDTA, pH 8.0) and incubated for 30 min at 37 °C. Afterwards, 2.5 µL of 10  mM dithiothreitol (DTT) were added and sample was incubated for 2  h at 37 °C. After that, 2.5 µL of 54  mM iodoacetamide (IAA) were added and sample was incubated for 1.5  h at room temperature, in the dark. Thus prepared protein was cleaned up using a MicroSpin G-25 column from GE Healthcare. The protein solution was diluted 1:1 (v/v) with 100  mM ammonium bicarbonate (NH4HCO3) and 500 ng of trypsin were added. The digestion was performed at 37 °C overnight. Finally, the sample was dried again and re-dissolved in 10 µL of 0.1% trifluoroacetic acid (TFA) for subsequent ZipTip C18 cleaning. Purified peptide solutions were dried and stored at 4 °C until analysis. Prior to analysis, peptides were redis-solved in 5 μL of α-cyano-4-hydroxycinnamic acid [CHCA, 6 mg mL−1, in 0.1% TFA/acetonitrile (50:50, v/v)] and two drops per 2 μL were spotted on the MALDI-MS target and dried at room temperature.

MALDI mass spectrometry

The MALDI-MS measurements were performed with a curved field reflectron time-of-flight (TOF) mass spec-trometer (AXIMA CFR+, Shimadzu Kratos Analytical, Manchester, UK) or with a TOF/RTOF mass spectrom-eter (AXIMA TOF2, Shimadzu Kratos Analytical). Both instruments are equipped with a pulsed nitrogen laser. The instruments were operated in the positive ion mode apply-ing an accelerating voltage of 20  keV. Delayed extrac-tion was used and the delay time was set according to the molecular mass of the analytes of interest in order to optimize mass spectrometric resolution. Typically, peptide

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mass fingerprint spectra were acquired by averaging 225 single, unselected laser shots. External calibration was per-formed using: the trimeric cluster of the MALDI matrix molecule (CHCA), bradykinin 1–5, bradykinin 1–7, angio-tensin II, angiotensin I, Glu1-Fibrinopeptide B, N-Acetyl renin, ACTH fragment 1–17, ACTH fragment 18–39, and ACTH fragment 7–38. Sequence tag information for up to five most prominent peaks (protonated molecules) from the acquired peptide mass fingerprints were obtained by means of post source decay (PSD) fragmentation and acquired by averaging 2250 laser shots. All mass spectra and MS/MS product ion spectra shown were smoothed using the com-pany-supplied Savitzky–Golay algorithm.

Protein identification

A database search for peptide mass fingerprint (PMF) and sequence tag data was performed against SwissProt 2015_8 and NCBInr 20150823 databases and Viridiplantae taxa using programs mMass (Strohalm et  al. 2010) and Mas-cot (http://www.matrixscience.com/). Fixed settings were: precursor ion mass tolerance ±0.5  Da, product ion mass tolerance ±1.0  Da, one missed trypsin cleavage site and carbamidomethylation of cysteines. Ammonia loss from N-terminal cysteine and oxidation of methionine were con-sidered as flexible modifications.

Statistical analysis

Presented results are average values ± standard deviation of at least three measurements. The results of each activ-ity assay were compared by analysis of variance (ANOVA), followed by Duncan test using the STATISTICA 12.0 (Stat Soft Inc., Tulsa, OK, USA) software package. Differences among means were considered statistically significant at p ≤ 0.05.

Results and discussion

It is crucial to establish an efficient protocol for separa-tion and purification of plant esterases in order to identify each isoform and ascertain its function. The complexity of numerous esterase isoforms that appear in each tissue (due to large number of potential esterase genes, complex regula-tion of protein expression, and numerous post-translational modifications of proteins), together with lack of sequence data make this task extremely challenging. Horizontal pla-nar IEF system used in this research enabled separation of 6 bands of esterases, within wide pI range (E1–E6, Fig. 1). Pronounced differences in expression levels among ana-lyzed tissues were observed for esterases with pI 9.3, 8.6 and 8.2 (E1, E2 and E3; respectively). Namely, isoesterases

E1 had higher staining intensity in L (plantlet leaves) tissue compared to TM and particularly TN, while E2 isoforms were more pronounced in both tumor tissues compared to L. Moreover, esterase band E3 showed enhanced in-gel activity in TN tissue compared to L and particularly TM. These results are in agreement with those of Krsnik-Rasol et al. (1999) previously obtained on a different IEF system.

In the study presented here, we set out to describe detected esterases from in vitro-grown horseradish tissues. We focused on finding a reliable protocol for detection, purification and identification of esterases from horseradish tissues. Since we expected the protein extracts to be com-plex mixtures to be analyzed as such, we decided to purify esterases as much as possible and then start the attempt of their identification.

The first step of esterase purification

In the first purification step, protein extracts from three types of horseradish tissue (leaf–L, teratoma–TM, tumor–TN) were subjected separately to fractionation on cation exchange chromatography on SP-Sepharose FF column. Results of this chromatography are presented in Fig. 2 for the sample of TN horseradish tissue line. Since L and TM samples gave very similar patterns to TN, these results are not shown. Protein-containing fractions were collected and tested for esterase activity (spectrophoto-metrically and in gel after IEF). Activity was spectrophoto-metrically measured separately towards 1- and 2-naphthyl

Fig. 1 Different isoesterase pattern in horseradish tissues of different morphology level. Horizontal planar isoelectric focusing (IEF) was used on gel with pH gradient 3–9. Marker proteins were stained with silver and proteins from horseradish tissue samples were visualized using esterase substrates (1- and 2-naphthyl acetate). P—pI marker proteins, L—leaf, TM—teratoma, TN—tumor. pIs of marker proteins are noted on the left, and pIs of isoesterases on the right

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acetate (1-NA and 2-NA, respectively, Table  1), as broad esterase substrates used in many plant studies (Balen et al. 2003; Radić and Pevalek-Kozlina 2010; Pavoković and Krsnik-Rasol 2012). Statistical analysis showed that pro-tein extracts from all three tissues had the same activity

towards 1-NA, while protein extract from leaves had higher activity towards 2-NA than extracts from other two tissues (Table  1). Regarding fractions collected after chromatog-raphy, the pattern of esterase activity was the same in all tissues extracts: already one purification step led to the

Fig. 2 Cation exchange chromatography of TN horseradish tissue line protein extract on SP-Sepharose FF. A chromatogram; a unbound on column, b eluted with 0.2 M NaCl, c eluted with 0.4 M NaCl, d eluted with 0.6  M NaCl, e eluted with 1.0  M NaCl. B SDS–PAGE of combined protein-containing fractions (silver staining) and IEF

of combined protein-containing fractions (visualization with ester-ase substrates, 1- and 2-naphthyl actetate); S—starting sample, P—molecular weight markers. Markers molecular weights (in kDa) are given on gel

Table 1 Activity of fractionated protein extracts from different horseradish tissues towards selected esterase substrates

* The activity values are expressed as means ± standard deviation of at least three measurements. If values are marked with different letters, it means they are significantly different (p ≤ 0.05) between different tis-sues within each fraction according to Duncan test1-NA 1-naphthyl acetate, 2-NA 2-naphthyl acetate. n.d. not determined (due to low protein amount and con-centration)

Starting tissue Fraction Protein concentration (mg mL−1)

Activity towards 1-NA(µmol min−1 mg−1)

Activity towards 2-NA(µmol min−1 mg−1)

L Protein extract 0.680 ± 0.059 0.351 ± 0.066a* 0.408 ± 0.054a

a 1.102 ± 0.158 0.306 ± 0.002b 0.267 ± 0.004b

b 1.249 ± 0.097 4.458 ± 0.102a 1.593 ± 0.025a

c 0.560 ± 0.000 0.780 ± 0.027a 0.256 ± 0.029a

d 0.021 ± 0.014 n.d.c n.d.c

e 0.018 ± 0.010 n.d.c n.d.c

TM Protein extract 1.646 ± 0.237 0.391 ± 0.029a 0.330 ± 0.038b

a 0.560 ± 0.053 0.421 ± 0.007a 0.379 ± 0.006a

b 0.651 ± 0.174 2.556 ± 0.061b 1.039 ± 0.023b

c 1.841 ± 0.231 0.374 ± 0.055b 0.230 ± 0.027a

d 0.267 ± 0.009 0.249 ± 0.040b 0.424 ± 0.047a

e 0.120 ± 0.032 0.163 ± 0.023b 0.000 ± 0.000b

TN Protein extract 2.025 ± 0.214 0.351 ± 0.055a 0.304 ± 0.032b

a 0.971 ± 0.232 0.281 ± 0.026b 0.256 ± 0.024b

b 0.392 ± 0.009 0.813 ± 0.091c 0.238 ± 0.024c

c 1.271 ± 0.169 0.278 ± 0.026c 0.161 ± 0.021b

d 0.529 ± 0.020 0.208 ± 0.022b 0.115 ± 0.018b

e 0.376 ± 0.055 1.155 ± 0.048a 0.501 ± 0.057a

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enrichment of the fraction ‘b’ (elution with 0.2 M NaCl in starting buffer) with NA-hydrolyzing esterases. This was particularly pronounced with the application of the 1-NA (specific activity increase of 2.3–12.7-fold, Table 1). Inter-estingly, in protein extract from tumor (TN), fraction ‘e’ also had heightened esterase activity, towards both sub-strates (Table 1). However, as the increase in specific activ-ity was so high for fraction ‘b’ in other two tissues extracts and considering the results of IEF (Fig.  2b), purification was continued on this fraction. Variation in sensitivity of plant esterases towards 1-NA and 2-NA was already reported for other plant species (Balen et  al. 2004; Radić and Pevalek-Kozlina 2010; Pavoković and Krsnik-Rasol 2012). Results obtained with spectrophotometric measure-ments were additionally confirmed by IEF, where the frac-tion ‘b’ showed enhanced staining of esterases, clearly vis-ible on the gel as dark bands (Fig. 2b).

The second step of esterase purification

The esterase patterns from all three horseradish tissues were almost identical after both IEF (although with differ-ent esterase expression levels) and fractionation by means of cation exchange chromatography on SP-Sepharose FF column (data not shown); therefore, we chose TN tissue for further analyses. Hence, after the first fractionation step on SP-Sepharose FF, several chromatographic procedures were applied.

The applied fast protein liquid chromatography (FPLC) methods led to separation of several protein peaks (data

not shown), but no method resulted in significant increase in esterase(s) purity (no significant increase in specific esterase activity was observed in any of collected frac-tions, as judged from spectrophotometric measurements and substrate staining of IEF gels). Re-chromatography of the fractions ‘b’ from SP-Sepharose FF on a smaller-sized SP-Sepharose FF column gave the best results in the terms of esterase purification (Fig.  3). Six protein peaks were separated by this type of chromatography and three of them were enriched in esterase activity (Fig.  3; Table 2). Statistical analysis was used in this case to com-pare resulting fractions amongst themselves. Clearly, the highest increase in specific activity for both substrates was in fractions 57–68 (9.7-fold for 1-NA and 5.4-fold for 2-NA). Following are the fractions 46–56 (2.2-fold increase in specific activity for 1-NA and 1.9-fold for 2-NA) and 122–128 (1.2-fold increase in specific activity for 1-NA and 2.5-fold for 2-NA). Interestingly, no activ-ity was observed after fractions 57–68 were resolved on IEF gel and stained with esterase substrates (Fig.  3b). The reason for this odd phenomenon lays in the number of esterases present in this sample. While SDS electro-phoresis resolves four protein bands for fractions 46–56 and only one major protein band for fractions 122–128 (Fig.  3b), many protein bands are present in the sample of fractions 57–68 (data not shown). This explains why no signal is present for this sample in IEF analysis (when separated in IEF, each protein has too low esterase activ-ity to be detected), and why this sample was not taken

Fig. 3 Re-chromatography of fraction ‘b’ from SP-Sepharose FF on the same chromatographic stationary phase. a chromatogram; after 16 mL of washing with starting buffer (50 mM phosphate, pH 7.0), elution was performed with 84 mL of linear NaCl gradient (0–0.3 M) and 25 mL of 1 M NaCl in starting buffer; protein peaks exhibiting an increase in esterase activity and analyzed by MS are marked with asterisk. b SDS–PAGE of combined protein-containing fractions

(silver staining) and IEF of combined protein-containing fractions (visualization with esterase substrates, 1- and 2-naphthyl acetate); P—molecular weight markers. Markers molecular weights (in kDa) are given on the left side of the gel. Isoelectric points of separated esterases are given on the right side of the gel. Samples on gels are denoted with fraction numbers

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into MS analysis (the resulting peptide mass fingerprints would be too complex for any protein identification).

Identification of esterases

Samples from re-chromatography on SP-Sepharose FF (fractions 46–56, eluted with NaCl gradient and fractions 122–128, eluted with 1 M NaCl, Fig. 3), although low in protein content, exhibited better resolved protein pattern with fewer protein bands on SDS gels than fractions from other second-step chromatographies. Therefore, those SP-Sepharose FF fractions were subjected to in-solution diges-tion with trypsin. This procedure resulted in abundant pep-tide mass fingerprints (Fig.  4). Peptide mass fingerprints were found to be more complex for the fractions 46–56, compared to the fractions 122–128; which is in accordance with the number of the protein bands detected on SDS gels of the respective fractions (Fig.  3b). The protonated mol-ecules representing peptides related to isolated proteins

with the highest intensity were subjected to PSD fragment ion analysis and the resulting sequence tag data were sub-mitted to SwissProt and NCBInr databases search using mMass and Mascot programs. Identified peptides are listed in Table 3.

According to the results of the SDS–PAGE, the frac-tions 46–56 contained four major proteins with molecular weights (MW) of 57.0, 49.4, 36.8 and 32.5 kDa (Fig. 3b); while according to the IEF gel, several esterases are pre-sent with pI values estimated to be 8.6, 8.2, 6.9 and 5.6 (Fig.  3b). Database search suggests that a signal at m/z 1710.08 ± 0.13 corresponds to a peptide highly homolo-gous to a Pectinesterase/pectinesterase inhibitor 18 from A. thaliana; moreover, the peptide detected at m/z of 1004.70 ± 0.08 corroborates this finding (Table  3). MW and pI data within experimental error confirm that a pro-tein similar to the esterase moiety part of Pectinesterase/pectinesterase inhibitor 18 from A. thaliana was indeed present in the fractions 46–56. From the isoesterase pattern

Table 2 Activity of esterases from TN horseradish tissue line protein extract partially purified on re-SP-Sepharose FF towards selected esterase substrates

*The activity values are expressed as means ± standard deviation of at least three measurements. If values are marked with different letters, it means they are significantly different (p ≤ 0.05) between different frac-tions according to Duncan test1-NA 1-naphthyl acetate, 2-NA 2-naphthyl acetate. n.d. not determined (due to low protein amount and con-centration)

Fraction number Protein concentration(mg mL−1)

Activity towards 1-NA(µmol min−1 mg−1)

Activity towards 2-NA(µmol min−1 mg−1)

Starting sample 0.426 ± 0.031 0.813 ± 0.091c* 0.238 ± 0.024d

5–14 0.476 ± 0.075 0.205 ± 0.062d 0.119 ± 0.024e

26–33 0.024 ± 0.014 n.d.e n.d.f

46–56 0.620 ± 0.086 1.767 ± 0.100b 0.455 ± 0.043c

57–68 0.923 ± 0.079 7.854 ± 1.180a 1.296 ± 0.099a

122–128 0.165 ± 0.005 0.993 ± 0.169c 0.603 ± 0.068b

Fig. 4 Peptide mass fingerprints of TN horseradish tissue line frac-tions after re-chromatography of fraction ‘b’ from SP-Sepharose FF on the same chromatographic stationary phase. a fractions 46–56. b

fractions 122–128. Asterisks mark peptides for which MS/MS (PSD fragment ion analysis) were performed and protein identification attempted

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of different horseradish tissues obtained by horizontal pla-nar IEF it seems that this protein is in the group marked as E2, which is present with the highest abundance in the TM tissue compared to TN and L (Fig. 1). Peptides correspond-ing to protonated molecule signals at m/z of 960.62 ± 0.07, 1185.75 ± 0.09 and 2287.35 ± 0.18 seem to be homologous to a Curculin-like (mannose-binding) lectin family protein from A. thaliana, although the peptide as protonated mol-ecule at m/z 960.62 ± 0.07 exhibits also significant homol-ogy with Serine carboxypeptidase-like protein 10 from A. thaliana. Both proteins are plausible also according to the SDS–PAGE based MW data (Fig. 3b).

From Table  3, it can be concluded that the fractions 122–128 contained an almost completely purified protein similar to GDSL esterase/lipase ESM1 from A. thaliana. Even though identification was achieved by matching only one (unique) peptide, data on protein MW and pI values estimated from SDS–PAGE and IEF gels (Fig. 3b) corrobo-rate this conclusion. In the isoesterase pattern of different horseradish tissues presented on Fig. 1 it is visible that the theoretical pI value of this protein corresponds to the one in group E3, which is present with higher abundance in TN than in TM and L tissues, thus indicating its role in plant tissue differentiation.

Physiological roles of identified esterases

Pectinesterase/pectinesterase inhibitor 18 from A. thaliana (UniProt code Q1JPL7) is really an interesting protein. It belongs to the group 2 of pectinmethyl esterases (PMEs), so-called pre-pro-proteins, where the pre region is the sig-nal peptide and the pro sequence is hypothesized to play roles in folding mechanism, in subcellular targeting as an intramolecular chaperone, in conformational folding of mature enzyme, or in acting as an autoinhibitor during transport through the endomembrane system (Markovič and Janeček 2004). Detailed protein sequence analysis of more than 100 PMEs revealed significant differences among PMEs of different origin (Markovič and Janeček 2004). The abundance of pectin-degrading enzymes at different growth stages in plants suggests that these enzymes play

important roles in development, dormancy breakage, seed and pollen germination, fruit ripening, etc. (Bou Daher and Braybrook 2015). The only plant PME 3D-structure known so far, that of a carrot enzyme (PDB code 1GQ8), shows that plant PMEs are aspartyl esterases belonging to the family of parallel β-helix proteins (Johansson et al. 2002). This family comprises four classes of pectin-degrading enzymes, including PMEs, that demethylesterify plant cell wall pectins.

In A. thaliana genome, 66 genes were predicted to code for PMEs (InterPro domain IPR000080, carbohydrate esterase family CE-8 according to CAZy classification sys-tem, Lombard et  al. 2014). The high number of putative PME sequences in the Arabidopsis genome suggests that these proteins could be involved in different developmental processes and/or act on different classes of pectins. Indeed, RT-PCR study of the relative expression of the 66 genes encoding putative A. thaliana PMEs throughout eight silique developmental stages demonstrated high diversity of expression patterns (Louvet et al. 2006).

Pectinesterase/pectinesterase inhibitor 18 from A. thali-ana (UniProt code Q1JPL7) has not been thoroughly inves-tigated. Louvet et  al. (2006) classified it in phylogenetic group 1 and expression group A (16 PMEs mainly highly or uniquely expressed in silique, particularly during late developmental phases). In another study, it is shown to be highly expressed in the presence of biotic stress (especially in the presence of Agrobacterium), but not in the pres-ence of abiotic stresses as observed in other PMEs (Pel-loux et al. 2007). This could be correlated with our finding that the esterase isoform E2, which probably corresponds to Pectinesterase/pectinesterase inhibitor 18, was found to be present with the highest abundance in the TM tissue obtained after transformation with A. tumefaciens. Also, a new moonlighting function of mature form of this par-ticular PME was identified in ribosome inactivation (inhibi-tion of the translation process by removing single adenine residue of the large rRNA) by De-la-Peña et  al. (2008). It is also important to note that this enzyme was inac-tive (both as a PME and as ribosome inactivating protein) when expressed in Escherichia coli, which stresses out the

Table 3 Identification of proteins from TN horseradish tissue line extract after in-solution tryptic digestion of re-SP-Sepharose FF fractions 46–56 and 122–128

Fractions Peptide [M + H]+ (m/z ± SD)

Identification MW (kDa)/pI

46–56 960.62 ± 0.07 Serine carboxypeptidase-like 10 or 47.4/6.2Curculin-like (mannose-binding) lectin family protein 49.1/7.8

1004.70 ± 0.08 Pectinesterase/pectinesterase inhibitor 18 58.1/8.61185.75 ± 0.09 Curculin-like (mannose-binding) lectin family protein 49.1/7.81710.08 ± 0.13 Pectinesterase/pectinesterase inhibitor 18 58.1/8.62287.35 ± 0.18 Curculin-like (mannose-binding) lectin family protein 49.1/7.8

122–128 1038.54 ± 0.11 GDSL esterase/lipase ESM1 41.0/8.1

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importance of post-translational modifications (e.g. glyco-sylation) for the function of eukaryotic enzymes (De-la-Peña et al. 2008). Clearly, an enzyme similar to Pectinester-ase/pectinesterase inhibitor 18 from A. thaliana would also have important role in horseradish defense system.

ESM1 (UniProt code Q9LJG3) belongs to the GDSL family of enzymes (InterPro domain IPR001087), which has 108 members in A. thaliana proteome. Bioinformatic analysis showed that the vast majority (99) of GDSL enzymes in A. thaliana contain signal peptides, indicating localization of these enzymes in extracellular parts or vari-ous organelles (Ling 2008). Suggested physiological roles mainly involve regulation of growth and development; including seed germination, flowering and defense reac-tions (Ling 2008).

Arabidopsis thaliana ESM1 was identified as Epithio-specifier modifier 1, which controls glucosinolate hydroly-sis (Zhang et al. 2006). Glucosinolates are sulfur-rich plant secondary metabolites whose basic skeleton consists of a β-thioglucose residue, an N-hydroxyiminosulfate moi-ety, and a variable side chain. They are not bioactive until enzymatically hydrolyzed, which occurs during tissue dis-ruption typically associated with a herbivore or pathogen. Horseradish TM and TN tissues, which were both obtained after cell transformation with A. tumefaciens, exhibited modified glucosinolate patterns compared to untransformed plantlet, namely L tissue type (Radojčić Redovniković et al. 2008). Hydrolysis of a single glucosinolate can produce a nitrile, an isothiocyanate, or a thiocyanate; with the final structure controlling the bioactivity (Lambrix et al. 2001). Functional A. thaliana ESM1 controls the ratio of nitrile to isothiocyanate production during glucosinolate hydrolysis, favoring production of toxic isothiocyanate, thus increasing plant defense against herbivores (Zhang et al. 2006). Also, it is possible that the remaining activation products could provide anti-microbial activity that would potentially steri-lize any cut site. Additionally, in the case of indole glucosi-nolates, this might yield a pool of indole-acetonitriles that could be utilized by the plant to make defensive compounds at the site of disruption (Burow et al. 2008). In the view of previously obtained results (Radojčić Redovniković et  al. 2008), presumably, a homologous protein in A. lapathifolia would have the same function(s).

Conclusions

Microbial esterases are already used in a wide range of biotechnological applications, whereas plant esterases are on the way to be exploited. Their potential has been wid-ened by tremendous progress in bioanalytical and protein engineering methods, which enable the production of con-siderable amounts of enzymes for further in-depth studies.

The striking example of plant esterase use in agriculture is a biosensor, i.e. a diagnostic tool, for organophosphate, neurotoxic insecticide, based on plant esterase-chitosan-gold nanoparticles-graphene (Bao et al. 2015). This result points up new possibilities of using esterase as biocontrol agent that can elicit defense reactions in plant cell and act at as a direct fungal inhibitor. Our research is focused on identification and characterization of horseradish esterases with the potential to be applied in biotechnology. Only par-tial purification (via two cation exchange chromatographic stages) of esterases from horseradish tumor and in-solution digestion by trypsin prior to MALDI-MS and MS/MS led to the identification of two probable esterase proteins. One is similar to GDSL esterase/lipase ESM1 from A. thaliana and seems to be upregulated in horseradish TN tumor. It has a molecular weight of ~43 kDa and isoelectric point at pH ~ 8.2, and is most likely involved in plant defense against herbivores. The second esterase is similar to Pectinester-ase/pectinesterase inhibitor 18, also from A. thaliana, and seems to be upregulated in both tumor tissues compared to leaf. This protein has a molecular weight of ~57.0 kDa and isoelectric point at pH ~ 8.6. It is also likely to be involved in plant defense, especially in presence of biotic stress. As plant esterases are involved in many important biological processes, we plan to continue our research on purification, biological function and structural characterization of ester-ases from different horseradish tissues.

Acknowledgements This work was supported by the Ministry of Science, Education and Sports of the Republic of Croatia, grant No. 098-1191344-2943 and 119-1191196-1200, and Bilateral Coopera-tion Grant Croatia-Austria (1/2010 to ILA and MMD).

Author contributions ILA performed purification of esterases, spectroscopy measurements and isoelectric focusing; participated in mass spectrometry measurements and wrote the manuscript. PPŠ per-formed plant tissue propagations and protein extractions and partici-pated in writing the manuscript. BB performed electrophoretic analy-ses and participated in writing the manuscript. MMD participated in designing and performing mass spectrometry measurements, as well as interpretation of the results. GA and BKP participated in planning the experiments, interpretation of the results and revising the manu-script. All authors read and approved the final manuscript.

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