Next Article in Journal
Mechanistic Insight into the 2° Alcohol Oxidation Mediated by an Efficient CuI/L-Proline-TEMPO Catalyst—A Density Functional Theory Study
Next Article in Special Issue
Catalytic Characteristics of New Antibacterials Based on Hexahistidine-Containing Organophosphorus Hydrolase
Previous Article in Journal
The Beneficial Sinergy of MW Irradiation and Ionic Liquids in Catalysis of Organic Reactions
Previous Article in Special Issue
Characterization of a Novel Nicotine Hydroxylase from Pseudomonas sp. ZZ-5 That Catalyzes the Conversion of 6-Hydroxy-3-Succinoylpyridine into 2,5-Dihydroxypyridine
Article Menu
Issue 9 (September) cover image

Export Article

Catalysts 2017, 7(9), 263; doi:10.3390/catal7090263

Article
Regioselective Synthesis of Lactulose Esters by Candida antarctica and Thermomyces lanuginosus Lipases
Luis F. Chávez-Flores 1, Hiram Isaac Beltran 2, Daniel Arrieta-Baez 3 and Dolores Reyes-Duarte 4,*
1
Posgrado en Ciencias Naturales e Ingeniería, Universidad Autónoma Metropolitana-Unidad Cuajimalpa, Av. Vasco de Quiroga 4871, Col. Santa Fe Cuajimalpa, Deleg. Cuajimalpa de Morelos 05348 CDMX, Mexico
2
Departamento de Ciencias Naturales, Universidad Autónoma Metropolitana-Unidad Cuajimalpa, Av. Vasco de Quiroga 4871, Col. Santa Fe Cuajimalpa, Deleg. Cuajimalpa de Morelos 05348 CDMX, Mexico
3
Instituto Politécnico Nacional - CNMN, Calle Luis Enrique Erro s/n, Unidad Profesional Adolfo López Mateos, Col. Zacatenco, México D.F. CP 07738, Mexico
4
Departamento de Procesos y Tecnología, Universidad Autónoma Metropolitana-Unidad Cuajimalpa, Av. Vasco de Quiroga 4871, Col. Santa Fe Cuajimalpa, Deleg. Cuajimalpa de Morelos 05348 CDMX, Mexico
*
Correspondence: Tel.: +52-55-581-465-00 (ext. 3872)
Received: 14 August 2017 / Accepted: 29 August 2017 / Published: 3 September 2017

Abstract

:
The interest in sugar esters as emulsifiers has been increasing in recent years because they have tunable surfactant properties that depend on the chain length of the fatty acid and the type of the sugar, covering a wide range of hydrophilic-lipophilic balance (HLB). In this work, ten biocatalysts were used for the transesterification reaction screening of lactulose, a prebiotic sugar, with vinyl laurate. The reactions were followed by thin layer chromatography (TLC) analysis, identifying two major monoesters mixtures defined as monoester fraction 1 and monoester fraction 2. Candida antarctica lipase B (Novozym 435) produces “monoester fraction 1”, while Thermomyces lanuginosus lipase (Lipozyme® TL IM) and Mucor miehei lipase (Lipozyme®) seem to produce the same “monoester fraction 2”. These three enzymes were selected as model biocatalysts for a kinetic study, and monoester fractions 1 and 2 from Novozym 435 and Lipozyme® TL IM, respectively, were used for product characterization. Monoester fraction 1 contained 86.9% of the major monoester in position 1-O-, and monoester fraction 2 contained 91.4% of 6′-O-. Although these lipases acylated three positions of lactulose, they mainly synthesize a monoester presenting regioselectivity. These results contribute to the study of the chemical structure diversity of biosurfactants to enhance their applications in foods, pharmaceutical products, and cosmetics.
Keywords:
biocatalysts; lipases; lactulose; prebiotic sugar; sugar fatty acid esters; transesterification; regioselectivity

1. Introduction

Sugar fatty acid esters have a wide range of applications in the food, cosmetic, detergent, and pharmaceutical industries [1]. They present interesting properties as surfactant [2], antimicrobial [3,4,5,6,7,8], antitumoral [9], and insecticide agents [10]. The degree of esterification and the type of fatty acid used determine their surfactant properties. It has also been demonstrated that sucrose esters (mono-, di-, or tri-esters) are hydrolysed to sucrose and fatty acids prior to intestinal absorption [11,12], giving them a GRAS (Generally Recognized As Safe) status [13]. Although different sugars and acyl donors have been widely studied [1,6,14,15,16,17], the use of a single sugar with prebiotic properties, such as lactulose, has been less studied [18,19] and might open significant applications in the food and pharmaceutical fields.
Lactulose is the simplest existent prebiotic. It is a disaccharide composed of galactose and fructose (4-O-β-d-galactopyranosyl-d-fructofuranose), and, in contrast to lactose, cannot be hydrolyzed due to its resistance to digestive enzymes, and therefore reaches the colon without alteration. There, lactulose is selectively metabolized by bifidobacterias and lactobacillus to lactic and acetic acids, reducing intestinal pH. These actions improve the gastrointestinal environment and microbiota presence and help to avoid the occurrence of constipation [20]. Therefore, lactulose is extensively used in infant and senior nutrition as a prebiotic [21].
On the other hand, lipases’ regioselectivity has been explored and used as an advantage for enzymatic synthesis over chemical synthesis. Nevertheless, due to the abundance of different hydroxyl groups in sugars, it is necessary to evaluate the behavior of enzymes to achieve a “directed enzymatic synthesis” for selective sugar esters. Enzymatic esterification using fructose, galactose-sugars that form lactulose-, lactose, and sucrose have been previously reported. Figure 1 shows the acylation positions of these sugars according to the following tendency: (a) fructose: 6-O- [22,23,24,25,26], 1-O- [23,27], and 1,6-O- [26]; (b) galactose: 6-O- [23,28,29]; (c) sucrose: 1′-O- [30,31,32], 6-O- [33,34,35,36,37], 6′-O- [33,34,35], 6,1-O-, and 6,6′-O- [37]; (d) lactose: 6′-O- [34]; (e) lactulose: 1-O- [30]. Other studies on the synthesis of lactulose esters have been previously reported; however, to the best of our knowledge, the acylation position has not been described [18,19].
In this work, we study the regioselective synthesis of lactulose esters, controlling the esterification position into the lactulose molecule as well as the esterification degree through modulating the biocatalyst used and the solvent composition in order to modify the structures of the lactulose esters and their properties.

2. Results

2.1. Enzymatic Synthesis of Lactulose Esters and Screening with Different Biocatalysts

The synthesis of lactulose esters was developed by transesterification with vinyl laurate, an acyl donor that shifts the equilibrium reaction towards ester formation due to a well-known vinyl alcohol tautomerization to acetaldehyde and product evaporation [37]. The enzymatic model reaction is shown in Figure 2.
Ten biocatalysts were used for the transesterification reaction screening, and they are shown in Figure 3a,b. Candida antarctica lipase B immobilized in acrylic resin (Novozym 435), Mucor miehei lipase immobilized on a macroporus ion-exchange resin (Lipozyme®), Thermomyces lanuginosus lipase immobilized as silica granulated (Lipozyme® TL IM), and Alcaligenes sp. lipase immobilized in granulated diatomaceous earth (QLG) showed activity and different products were observed. The monoesters produced with Lipozyme TL IM, Lipozyme, and QLG have the same retention factor (Rf = 0.289), while with Novozym 435 the Rf is 0.235.
Three biocatalysts were tested by their ability to synthesize lactulose esters. After one hour, the reaction products were followed by TLC (Figure 4). As observed here, there are two major groups of monoesters, defined as monoester fraction 1 and monoester fraction 2. The Novozym 435 biocatalyst produces “monoester fraction 1”, while Lipozyme TL IM and Lipozyme from Mucor miehei lipases seem to produce the same “monoester fraction 2”. These three enzymes were selected as model biocatalysts for the kinetic study, and monoester fractions 1 and 2 from the Novozym 435 and Lipozyme TL IM lipases were used for product characterization.
The monoester fractions obtained were purified by liquid–liquid extraction and flash chromatography, and were characterized by 1H, 13C, and two-dimensional (2D) (HMBC) NMR, MS-ESI, and FT-IR.

2.2. Characterization by FT-IR, 1H, 13C, and 2D (HMBC) NMR and MS-ESI

FT-IR: The analysis of the FT-IR data has shown a specific band at 1740 cm−1, clearly stating the presence of C=O ester stretching vibrations.
Electrospray Mass Spectrometry (MS-ESI) Analysis: The products’ reaction was first analyzed by MS-ESI (+). Electrospray is a technique where droplets are generated when a high voltage is applied to a liquid stream. In the ESI process, a population of variably charged ions is generated. In this study, the monoesters’ spectra mainly contain [M + H]+, [M + Na]+, or [M + K]+.
Both mixture reactions were injected separately in a direct form into the spectrometer to analyze the products formed. According to the ESI (+) mass spectra, both enzymes produce a monoester with an m/z of 547.2738 (For a [C24H44O12 + Na] + calc., m/z 547.2730). However, when enzyme Lipozyme TL IM was used instead of Novozym 435, the reaction products were purer (Figure 5).
The signals observed in the products from Lipozyme TL IM belong to the monoester fragmentation, as we can see in Figure 6b, and this fragmentation was not observed when products from Novozym 435 were analyzed (Figure 6a), indicating that it is not the same product.
NMR Analysis: The 1H-NMR spectrum of the monoester fractions from the lipase-catalyzed reactions clearly shows a group of signals that belong to more than one product, which means that there are at least two different acylation positions in monoesters (Table 1).
According to the 13C NMR analysis, three different lactulose monoesters could be identified. When the acylation occurs in position 1 of the fructose moiety (Monoester-1 (1-O-), Table 1) the chemical shift of the C-1 was found at 64.0 ppm, whereas if the acylation is located in position C-6′ of the galactose moiety the signal is located at δ 62.63 ppm (Monoester-6′ (6′-O-), Table 1). These observations corroborated with the predicted carbonyl signals of the monoesters at the 13C NMR. Two carbonyl signals were observed at δ 173.5 ppm for the acylation in position C-1 of the fructose, and another signal at δ 174.1 ppm for the acylation in position C-6′ of the galactose. The NMR analysis of the fractions showed two signals that could indicate the presence of another monoester. Signals at δ 64.21 and 173.8 ppm belong to another structure, probably to a monoester with acylation in position C-6 of the fructose moiety (Monoester-6 (6-O-), Table 1). This signal showed correlation in HMBC indicating the presence of this monoester. The 1H and 13C NMR spectra can be viewed in Supplementary Figures S1a,b and S2a,b.
Signals assignments where corroborated with the HMBC 2D NMR spectral analysis of the products by the correlation of the carbonyl signals with the C-2″ of the lauroyl chain (Figure S3a,b).
According to the observations in the 13C NMR spectrum (Figure S2a), three products (Table 1) were found in the Novozyme 435-catalyzed reaction. However, only two monoesters (1-O- and 6′-O-) were identified when Lipozyme TL IM was used to catalyze the reaction (Figure S2b) (Table 1). This could be explained from the point of view of the steric hindrance to achieve ester formation. As we could see in Figure 7, -OH in position 6′ of the galactose moiety are more sterically available instead of C-OH in position 6 of the fructose moiety. Therefore, this means that the lipase Lipozyme TL IM is more selective to produce only two isomers of the monoester, while the lipase Novozym 435 does not have this restriction to produce more isomers, as it was demonstrated with this analysis.
The proportion of monoesters synthesized by lipases from Novozym 435 and Lipozyme TL IM was determined by a 1H NMR signal integration of the characteristic chemical shifts at each monoester, where the characteristic chemical shifts of each molecule were identified. This was also performed in order to aid in the quantification of each molecule, and this was feasible because each molecule presented a characteristic chemical shift without overlap among at least one of their signals and thus were differentiable to determine proportions through signal integration.
As stated, the 1H NMR signals corresponding to each monoester were integrated to identify and quantify the studied molecules, and each one appeared as a doublet with displacements of 4.39 (Monoester-1 (1-O)), 4.42 (Monoester-6′ (6′-O)), and 4.45 (Monoester-6 (6-O)) ppm from Novozym 435 (Figure S4) and from Lipozyme TL IM at 4.39 (Monoester-1 (1-O)) and 4.42 (Monoester-6′ (6′-O)) ppm (Figure S5).
The signals from each monoester contained in the mixture generated by Novozym 435 were identified and integrated: a doublet to -H of C-1 4.39 ppm and integration of 1, -H of C-6′ 4.42 ppm and integration of 0.1, and -H of C-6 at 4.45 ppm and integration of 0.05. The integration of the proton related to C-1′ appearing as a doublet at 3.92 ppm resulted in 1.15 (Figure S4), coinciding with the sum of the integrals of the three doublets described above, and also indicating that no other isomers or related structures were present. The mixture contained 87% of 1-O-, which is the major monoester, 8.7% of 6-O-, and 4.3% of 6′-O-.
The signals in the 1H spectra resulting from the Lipozyme TL IM catalyzed-synthesis were identified: 3.92 ppm, 3.93 ppm, and integration of 1.09 for the doublet proton related to C-1′, -H of C-1 at 4.39 ppm and integration of 0.093, and -H of C-6′ at 4.42 ppm and integration of 1 (Figure S5). The mixture contained 91.5% of the major monoester (6′-O-) and 8.5% of the minor one (1-O-). Although lipases from Novozym 435 and Lipozyme TL IM are able to acylate in different positions of lactulose, these mainly synthesized one monoester, presenting regioselectivity. Comparing the lipase acylation positions for disaccharides reported in the literature (Figure 1c,d), the produced monoesters also gathered the acyl moiety in the two less steric-hindered 1-O- and 6′-O- positions of lactulose, while the 6-O- position remained almost untouched, as occurred in the 1′-O- of sucrose. In this work, lipase B from Candida antarctica acylates the less steric-hindered position 1-O- with a yield of 86.9%. On the other hand, Thermomyces lanuginosus lipase prefers the acylation of lactose (Figure 1d) in position 6′-O- (in galactose), coinciding with this work where Lipozyme TL IM acyls the same position of lactulose (6′-O-) with a yield of 91.4%.

2.3. Conversion to Monoester by Selected Enzymes and Effect of Temperature in the Reaction

Figure 8 shows the time-course of the mixture of monoesters produced by each catalyzed reaction at 30 and 60 °C, observed as one monoester peak in a HPLC analysis. At 60 °C, Novozyme 435 and Lipozyme TL IM had similar behaviour: maximum monoesters production in short time (21.6 mm of monoesters in 1.5 h and 12.3 mm in 1 h, respectively). The decay in the concentration was due to the formation of diesters and triesters, observed by HPLC analysis (data not shown). Lipozyme from Mucor miehei achieved 21.6 mm in 5 h, with less conversion to diesters.
When the reaction temperature decreased from 60 to 30 °C, the synthesis of monoesters was preferred over diesters synthesis, and the effect could be translated into an increase in selectivity with a lower temperature. This effect could be observed with Novozyme 435 and Lipozyme TL IM but not with the Lipozyme from the Mucor miehei lipase. In the last case, when the temperature decreased, the synthesis of lactulose monoesters was maintained almost equal from 21.6 mm in 5 h to 20.9 mm in 7 h, and a lower diester production was also observed, possibly due to the active site structure of this type of lipase [38,39], as commented on in the next section.

Effect of Substrates Concentrations on Monoesters Yield

In order to evaluate the effect of the molar ratio sugar/acyl donor on the transesterification reaction, two conditions were evaluated (Figure 9). A ratio of 1:5 was better than 1:1 for achieving higher yields: Novozym 73% vs. 6% in 1.5 h; Lipozyme TL IM 43% vs. 23% in 1 h; Lipozyme from Mucor miehei 73% vs. 37% in 5 h. Two factors could be involved in this behaviour: (a) due to the lower concentration of acyl donor in the 1:1 reaction, the low water content present was enough for the vinyl laurate’s hydrolysis, and therefore the transesterification reaction to sugar esters decreased [40]; (b) the lactulose monoesters produced, due to their amphiphilic behaviour, could help to avoid sugar precipitation, stabilizing the reaction mixture and displacing the equilibrium of the reaction towards transesterification [41,42].

2.4. Calculation of the Hydrophilic-Lipophilic Balance (HLB)

The HLB of an emulsifier is an expression of its Hydrophile-Lipophile Balance, i.e., the balance of the size and strength of the hydrophilic (or polar) and the lipophilic (or nonpolar) groups of the emulsifier. All emulsifiers consist of a molecule that combines both hydrophilic and lipophilic groups [43].
The classification of surfactants by HLB numbers allows for the making of certain predictions of the behaviour of these compounds, facilitating the optimum selection of emulsifiers, wetting agents, and other types of agents, and making a connection between theory and practical applications [44].
According to the structures studied (Figure 2) and the Equations (1) and (3) (Davies method) and (2) (Griffin method) (Subsection 4.2.5), HLB values were calculated.

2.4.1. Griffin Method

  • Mh = 341.29 g/mol
  • MT-mono = 524.60 g/mol
  • MT-di = 706.91 g/mol
  • MT-tri = 889.22 g/mol
  • HLB mono = 20 (341.29/524.60) = 13.01 ≈ 13
  • HLB di = 20 (341.29/706.91) = 9.65 ≈ 10
  • HLB tri = 20 (341.29/889.22) = 7.67 ≈ 8
where:
  • HLBmono: HLB value by monoester
  • HLBdi: HLB value by diester
  • HLBtri: HLB value by triester

2.4.2. Davies Method

H L B = n ( # group   - O - C = O ) + n ( # group   - O - ) + n ( # group - OH ) n ( # group   - CH - ,   - CH 2 - ,   - CH 3 ,   - C - ) + 7
HLBmono =   ( 2.4 ) + 3 ( 1.3 ) + 7 ( 1.9 ) 23 ( 0.475 ) + 7 = 15.675 16
HLBdi =   2 ( 2.4 ) + 3 ( 1.3 ) + 6 ( 1.9 ) 34 ( 0.475 ) + 7 = 10.95 11
HLBtri =   3 ( 2.4 ) + 3 ( 1.3 ) + 5 ( 1.9 ) 45 ( 0.475 ) + 7 = 6.225 6
HLB scale goes from 1 to 20, where 1 is the most hydrophobic and 20 the most hydrophilic compound. The HLB values from the sucrose and lactulose esters are similar because of their chemical structures. According to their values, these sugar monoesters (Griffin 13 and Davies 16) could be used for dairy drinks, ice creams, or in the production of wheat products and detergents; the diesters (Griffin 10 and Davies 11) in the manufacture of chewing gum and wheat products; and the triesters (Griffin 8 and Davies 6) for processing milk powder and confectionery [2,45].

3. Discussion

Novozym 435 recognizes three acylation positions for lactulose monoesters (8.6% of 6-O- and 4.5% of 6′-O-) but shows preferred regioselectivity for 1-O- (86.9%), whereas Lipozyme TL IM shows regioselectivity for the less steric-hindered 1-O- (8.5%) and 6′-O- (91.4%) positions of lactulose (Figure 10).
The difference of this preference could be based on the structural and physicochemical properties of the active sites of these enzymes; whereas Thermomyces lanuginosus lipase (Lipozyme TL IM) has a large cleavage for attachment to alcohols and a narrow cleavage for acyl bonding, the lipase B of Candida antarctica (Novozym 435) has a large cleavage for attachment to the acyl and a small cleavage for attachment to alcohols [46]. This allows Candida antarctica lipase B to accept a wide type of large substrates, while Thermomyces lanuginosus has steric restrictions in its active site, only catalyzing on the most exposed acyl positions. These results match with the already known sn-1(3) specificity of the Thermomyces lanuginosus and Mucor miehei lipases and the non-regiospecificity of Candida antarctica lipase B over triglycerides [38,39]. This work supports the view that lactulose monoesters can be produced while controlling the molecular properties of each enzyme, and that the structure of the substrates and the parameters of the reaction affect the binding enzyme substrate [41]. As shown in this work and coinciding with previous studies, the use of a polar co-solvent (20% DMF) allows the reactions to produce more monoesters due to the improvement of the solubility of the sugar, producing the monoester as the main product [40].
On the other hand, Novozym 435 (reaction conditions: 30 °C and ratio sugar/acyl donor 1:5) is the best enzyme for the regioselective synthesis of the monoester 1-O- and Lipozyme TL IM for the monoester 6′-O- (reaction conditions: 60 °C and ratio sugar/acyl donor 1:5). Lipozyme from Mucor miehei presumably catalyzed the synthesis of the same monoesters as Lipozyme TL IM; in such a case, it would give higher yields than the last one for producing 6′-O-lactulose monoester (reaction conditions: 60 °C and ratio sugar/acyl donor 1:5). This work gives us the opportunity to produce new compounds with potential functional properties that have to be explored.

4. Materials and Methods

4.1. Materials

Lipozyme TL IM® and Novozym 435® were kindly donated from Novo Nordisk (Mexico City, Mexico); QLG (from Alcaligenes sp. from Meito Sangyo Co.( Tokyo, Japan)) was donated by F. Plou; and LEW-ROL-PEI and LEW-ROL-160 from Rhyzomucor oryzae immobilized in celite mono- and multi-point were donated by F.Valero (ENZNUT-BIOCATEM network). The following enzymes are from Sigma-Aldrich: Lipozyme® from Mucor miehei, lipase from porcine pancreas type II (lyophilized), Amano lipase PS from Burkholderia cepacia, Palatase® lipase from Rhizomucor miehei, and protease from Bacillus licheniformis.
Lactulose ≥95%, vinyl laurate, and 2-methyl-2-butanol (2M2B) were obtained from Sigma-Aldrich (St. Louis, MO, USA). Solvents such as dimethylformamide (DMF), chloroform, methanol, ethyl acetate, and acetonitrile were purchased from Fermont (Monterrey, México). Flash silica (gel 60, 230–400 mesh) was from Merck (Edo. de México, México). HPLC columns C-8 250 × 4.6 mm Microsorb-MV were from Varian. Chromatographic plates Polygram Sil G/UV254 (0.2 mm) were from Machery-Nagel (Düren, Germany).

4.2. Methods

4.2.1. Enzymatic Reactions

Lactulose laurate was synthesized by the transesterification of vinyl laurate (150 mm) and lactulose (30 mm) in 2M2B or tert-butanol with 20% DMF, with 25 mg/mL of biocatalysts, in a 5 mL scale. The reactions were performed at 30 and 60 °C with orbital shaking (600 rpm). The reactions were followed by TLC and HPLC. Different biocatalysts were used for the reaction screening using 25 mg/mL.

4.2.2. Products Purification

The synthesis was scaled up to a volume of 120 mL and incubated for one hour. Immediately after reaching the maximum yield for monoesters, the reaction was stopped to avoid the occurrence of diesters (determined by TLC and HPLC). Subsequently, the mixture was cooled and filtered. Purification continued in two steps: (i) a solvent extraction/partition using chloroform: hexane: acetonitrile (1:3.5:4.5 v/v), and (ii) flash chromatography on silica flash (Gel 40/140 Mesh) using ethyl acetate and methanol (9:1 v/v) as eluent.

4.2.3. TLC and HPLC Analysis

Analytical TLC was performed on silica gel plates using chloroform/methanol 4:1 (v/v) as eluent. Spots were revealed with a ceric ammonium sulfate staining solution and heating at 120 °C for 10 min.
The HPLC analysis was carried out by reverse-phase high-performance liquid chromatography (HPLC) using a Varian Prostar (Agilent, Santa Clara, CA, USA), a Microsorb-MV C18 column (250 × 4.6 mm) (Agilent, Santa Clara, CA, USA ), and an evaporative light-scattering detector (PL-ELSD 2100 Ice) (Polymer Laboratories-Agilent, Santa Clara, CA, USA) equilibrated at 60 °C The mobile phase initiated with acetonitrile:water 40:60 (v/v) at 1.2 mL/min for 4 min. Then, a gradient to 95% of acetonitrile was performed in 3 min and continued to pure acetonitrile for 20 min more. Acetonitrile was then held as a mobile phase for 7 min. Integration was carried out using the Galaxie software Varian (Agilent, Santa Clara, CA, USA). Calibration analyses were performed using monoesters obtained and purified as described above.

4.2.4. Products Characterization by FT-IR, 1H, 13C, and 2D NMR and MS-ESI

The FT-IR analysis was measured in a Bruker Tensor 27 FT-IR (Bruker Daltonics, Billerica, MA, USA). The mass spectrometry analysis was done using 1 mg of reaction products dissolved in 1 mL of methanol and directly injected into a Bruker MicrOTOF-QII system by an electrospray ionization (ESI) interface (Bruker Daltonics, Billerica, MA, USA) for its mass spectrometric (MS) analysis. The conditions of MS analysis in the positive ion mode were as follows: drying gas (nitrogen), flow rate, 8 L/min; gas temperature, 180 °C; scan range, 50–3000 m/z; end plate off set voltage, 500 V; capillary voltage, 4500 V; nebulizer pressure, 2.5 bar. The accurate mass data of the molecular ions were processed through the software DataAnalysis 4.0 (BrukerDaltonics, Billerica, MA, USA), which provided a list of possible elemental formulas using Generate Molecular Formula Editor, as well as a sophisticated comparison of the theoretical with the measured isotope pattern (σ value) for increased confidence in the suggested molecular formula (Bruker Daltonics Technical Note 008, 2004). The widely accepted accuracy threshold for confirmation of elemental compositions was established at 5 ppm. The instrument calibration was performed using a 74,900-00-05 Cole Palmer syringe pump (Cole Palmer, Vernon Hills, IL, USA) directly connected to the interface, with a sodium formate cluster solution.
The monoesters obtained from the lipase-catalyzed reactions were characterized by 1H- and 13C-NMR (AVANCE III, Bruker NMR System, 750 MHz) (BrukerDaltonics, Billerica, MA, USA). The NMR spectra were recorded in deuterated methanol (CH3OD). Chemical shifts (δ values) were given in ppm. ACD/CNMR Predictor 2015 (ACD-Labs, Toronto, Canada) was used to calculate the chemical shifts of the monoesters, using CH3OD as a solvent and 187.5 MHz as a theoretical frequency.

4.2.5. Calculation of HLB Values

The term HLB (hydrophilic-lipophilic balance) is a systematic scheme of emulsifiers suitable for any application. It is determined by two formulas. In the Griffin method, the HLB value of a nonionic surfactant is proportional to the weight of the fraction of the hydrophilic part: [47,48,49].
H L B = 20 M h M T
where:
  • Mh = weight fraction of the hydrophilic group
  • MT = the molecular weight of the whole molecule
The Davies method consists in the assumption that hydrophilic groups have a positive contribution to HLB number, while the hydrophobic groups have a negative effect. Davies calculated these contributions and named them as HLB group numbers.
Knowing the molecular structure, these values are added according to the following formula [50]:
H L B = n u m b e r   o f   h y d r o p h i l i c   g r o u p + n u m b e r   o f   l i p o p h i l i c   g r o u p + 7 .

5. Conclusion

In this work, we used lactulose for the synthesis of sugar esters. Under the studied conditions, using 2M-2B with DMF (20%), Novozym 435 preferably synthesized 1-O-lactulose monoester, whereas Lipozyme TL IM showed preferred regioselectivity for 6′-O-lactulose monoester. The products reached conversions over 60% (1:5, sugar:acyl donor) in less than 2 h. The conversions with lactulose were higher than with other disaccharides, like sucrose, as shown in previous reports. In this research, by using a prebiotic sugar, we demonstrated that it is possible to control the esterification degree in sugar ester synthesis through the modulation of the solvent composition and the biocatalyst, in order to obtain products with new potential properties, like prebiotic surfactants of interest in the food industry.

Supplementary Materials

The following are available online at www.mdpi.com/2073-4344/7/9/263/s1, Figure S1: 1H NMR spectra of purified reaction products: (a) Monoester fraction 1 of Novozym 435 and (b) Monoester fraction 2 of Lipozyme TL IM, Figure S2: 13C NMR spectra of purified reaction products: (a) Monoester fraction 1 of Novozym 435 and (b) Monoester fraction 2 of Lipozyme TL IM, Figure S3: HMBC spectra of purified reaction products: (a) Monoester fraction 1 of Novozym 435 and (b) Monoester fraction 2 of Lipozyme TL IM. X axis: 1H; Y axis: C., Figure S4: 1H NMR spectra and integration of monoesters obtained from Novozym 435, Figure S5: 1H NMR spectra and integration of monoesters obtained from Lipozyme TL IM.

Acknowledgments

This work has been supported by project CB-2015-258385 of the Consejo Nacional de Ciencia y Tecnología (CONACyT). L.F.C.F expresses gratitude for the PhD scholarship 372441 to CONACyT. We thank Dilyn Myers (Fulbright U.S. Program at UAM Cuajimalpa, 2016) for reviewing English text.

Author Contributions

D.R.-D conceived the concept. L.F.C.-F and D.R.-D designed and performed the experiments. D.A.-B and H.I.B contributed to the spectroscopic analysis. All of the authors contributed to analyzing the data and writing the manuscript. All of the authors have read and approved the final manuscript.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Ballesteros, A.; Plou, F.J.; Alcalde, M.; Ferrer, M.; García-Arellano, H.; Reyes-Duarte, D.; Ghazi, I. Enzymatic synthesis of sugar esters and oligosaccharides from renewable resources. In Biocatalysis in the Pharmaceutical and Biotechnology Industries; CRC Press: Boca Raton, FL, USA, 2006. [Google Scholar]
  2. Mitsubishi-Kagaku-Foods Corporation. Ryoto sugar esters technical information. 2004. Available online: http://www.mfc.co.jp/english/infor.htm (accessed on 3 September 2017).
  3. Abdulmalek, E.; Mohd Saupi, H.S.; Tejo, B.A.; Basri, M.; Salleh, A.B.; Raja Abd Rahman, R.N.Z.; Abdul Rahman, M.B. Improved enzymatic galactose oleate ester synthesis in ionic liquids. J. Mol. Catal. B Enzym. 2012, 76, 37–43. [Google Scholar] [CrossRef]
  4. Nobmann, P.; Smith, A.; Dunne, J.; Henehan, G.; Bourke, P. The antimicrobial efficacy and structure activity relationship of novel carbohydrate fatty acid derivatives against Listeria spp. and food spoilage microorganisms. Int. J. Food Microbiol. 2009, 128, 440–445. [Google Scholar] [CrossRef] [PubMed]
  5. Zhao, L.; Zhang, H.; Hao, T.; Li, S. In vitro antibacterial activities and mechanism of sugar fatty acid esters against five food-related bacteria. Food Chem. 2015, 187, 370–377. [Google Scholar] [CrossRef] [PubMed]
  6. Habulin, M.; Šabeder, S.; Knez, Ž. Enzymatic synthesis of sugar fatty acid esters in organic solvent and in supercritical carbon dioxide and their antimicrobial activity. J. Supercrit. Fluids 2008, 45, 338–345. [Google Scholar] [CrossRef]
  7. Devulapalle, K.S.; Gómez de Segura, A.; Ferrer, M.; Alcalde, M.; Mooser, G.; Plou, F.J. Effect of carbohydrate fatty acid esters on Streptococcus sobrinus and glucosyltransferase activity. Carbohydr. Res. 2004, 339, 1029–1034. [Google Scholar] [CrossRef] [PubMed]
  8. Karlová, T.; Polakova, L.; Šmidrkal, J.; Filip, V. Antimicrobial effects of fatty acid fructose esters. Czech J. Food Sci. 2010, 28, 146–149. [Google Scholar]
  9. Ferrer, M.; Soliveri, J.; Plou, F.J.; López-Cortés, N.; Reyes-Duarte, D.; Christensen, M.; Copa-Patiño, J.L.; Ballesteros, A. Synthesis of sugar esters in solvent mixtures by lipases from Thermomyces lanuginosus and Candida antarctica B, and their antimicrobial properties. Enzym. Microb. Technol. 2005, 36, 391–398. [Google Scholar] [CrossRef]
  10. Puterka, G.J.; Farone, W.; Palmer, T.; Barrington, A. Structure-function relationships affecting the insecticidal and miticidal activity of sugar esters. J. Econ. Entomol. 2003, 96, 636–644. [Google Scholar] [CrossRef] [PubMed]
  11. Noker, P.E.; Lin, T.H.; Hill, D.L.; Shigeoka, T. Metabolism of 14c-labelled sucrose esters of stearic acid in rats. Food Chem. Toxicol. 1997, 35, 589–595. [Google Scholar] [CrossRef]
  12. Shigeoka, T.; Izawa, O.; Kitazawa, K.; Yamauchi, F.; Murata, T. Studies on the metabolic fate of sucrose esters in rats. Food Chem. Toxicol. 1984, 22, 409–414. [Google Scholar] [CrossRef]
  13. Eldred, J.S. Gras Notification for Sucrose Fatty Acid Esters; Center for Food Safety and Applied Nutrition, Food and Drug Administration: Washington, DC, USA, 2003.
  14. Sutili, F.K.; Ruela, H.S.; Leite, S.G.F.; Miranda, L.S.d.M.; Leal, I.C.R.; de Souza, R.O.M.A. Lipase-catalyzed esterification of steric hindered fructose derivative by continuous flow and batch conditions. J. Mol. Catal. B Enzym. 2013, 85–86, 37–42. [Google Scholar] [CrossRef]
  15. Van Kempen, S.E.H.J.; Boeriu, C.G.; Schols, H.A.; de Waard, P.; van der Linden, E.; Sagis, L.M.C. Novel surface-active oligofructose fatty acid mono-esters by enzymatic esterification. Food Chem. 2013, 138, 1884–1891. [Google Scholar] [CrossRef] [PubMed]
  16. Van Kempen, S.E.H.J.; Schols, H.A.; van der Linden, E.; Sagis, L.M.C. Effect of variations in the fatty acid chain on functional properties of oligofructose fatty acid esters. Food Hydrocoll. 2014, 40, 22–29. [Google Scholar] [CrossRef]
  17. Horchani, H.; Aissa, I.; Ouertani, S.; Zarai, Z.; Gargouri, Y.; Sayari, A. Staphylococcal lipases: Biotechnological applications. J. Mol. Catal. B Enzym. 2012, 76, 125–132. [Google Scholar] [CrossRef]
  18. Bernal, C.; Illanes, A.; Wilson, L. Heterofunctional hydrophilic-hydrophobic porous silica as support for multipoint covalent immobilization of lipases: Application to lactulose palmitate synthesis. Langmuir 2014, 30, 3557–3566. [Google Scholar] [CrossRef] [PubMed]
  19. Bernal, C.; Illanes, A.; Wilson, L. Improvement of efficiency in the enzymatic synthesis of lactulose palmitate. J. Agric. Food Chem. 2015, 63, 3716–3724. [Google Scholar] [CrossRef] [PubMed]
  20. Corzo, N.; Alonso, J.L.; Azpiroz, F.; Calvo, M.A.; Cirici, M.; Leis, R.; Lombo, F.; Mateos-Aparicio, I.; Plou, F.J.; Ruas-Madiedo, P.; et al. [prebiotics: Concept, properties and beneficial effects]. Nutr. Hosp. 2015, 31 (Suppl. 1), 99–118. [Google Scholar] [PubMed]
  21. Panesar, P.S.; Kumari, S. Lactulose: Production, purification and potential applications. Biotechnol. Adv. 2011, 29, 940–948. [Google Scholar] [CrossRef] [PubMed]
  22. Lee, K.P.; Kim, H.K. Antibacterial effect of fructose laurate synthesized by Candida antarctica b lipase-mediated transesterification. J. Microbiol. Biotechnol. 2016, 26, 1579–1585. [Google Scholar] [CrossRef] [PubMed]
  23. Schlotterbeck, A.; Lang, S.; Wray, V.; Wagner, F. Lipase-catalyzed monoacylation of fructose. Biotechnol. Lett. 1993, 15, 61–64. [Google Scholar] [CrossRef]
  24. Scheckermann, C.; Schlotterbeck, A.; Schmidt, M.; Wray, V.; Lang, S. Enzymatic monoacylation of fructose by two procedures. Enzym. Microb. Technol. 1995, 17, 157–162. [Google Scholar] [CrossRef]
  25. Li, L.; Ji, F.; Wang, J.; Jiang, B.; Li, Y.; Bao, Y. Efficient mono-acylation of fructose by lipase-catalyzed esterification in ionic liquid co-solvents. Carbohydr. Res. 2015, 416, 51–58. [Google Scholar] [CrossRef] [PubMed]
  26. Li, L.; Ji, F.; Wang, J.; Li, Y.; Bao, Y. Esterification degree of fructose laurate exerted by Candida antarctica lipase b in organic solvents. Enzym. Microb. Technol. 2015, 69, 46–53. [Google Scholar] [CrossRef] [PubMed]
  27. Tükel, S.; Sahin, P.B.; Yildirim, D. Optimization of lipase-catalyzed synthesis of fructose stearate using response surface methodology. Artif. Cells Nanomedicine Biotechnol. 2013, 41, 344–351. [Google Scholar] [CrossRef] [PubMed]
  28. Chen, J.; Kimura, Y.; Adachi, S. Continuous synthesis of 6-O-linoleoyl hexose using a packed-bed reactor system with immobilized lipase. Biochem. Eng. J. 2005, 22, 145–149. [Google Scholar] [CrossRef]
  29. Kitagawa, M.; Fan, H.; Raku, T.; Shibatani, S.; Maekawa, Y.; Hiraguri, Y.; Kurane, R.; Tokiwa, Y. Selective enzymatic preparation of vinyl sugar esters using dmso as a denaturing co-solvent. Biotechnol. Lett. 1999, 21, 355–359. [Google Scholar] [CrossRef]
  30. Riva, S.; Nonini, M.; Ottolina, G.; Danieli, B. Subtilisin-catalyzed esterification of di-and oligosaccharides containing a d-fructose moiety. Carbohydr. Res. 1998, 314, 259–266. [Google Scholar] [CrossRef]
  31. Potier, P.; Bouchu, A.; Gagnaire, J.; Queneau, Y. Proteinase N-catalysed regioselective esterification of sucrose and other mono- and disaccharides. Tetrahedron Asymmetry 2001, 12, 2409–2419. [Google Scholar] [CrossRef]
  32. Wang, X.; Miao, S.; Wang, P.; Zhang, S. Highly efficient synthesis of sucrose monolaurate by alkaline protease protex 6l. Bioresour. Technol. 2012, 109, 7–12. [Google Scholar] [CrossRef] [PubMed]
  33. Woudenberg-van Oosterom, M.; van Rantwijk, F.; Sheldon, R.A. Regioselective acylation of disaccharides in tert-butyl alcohol catalyzed by Candida antarctica lipase. Biotechnol. Bioeng. 1996, 49, 328–333. [Google Scholar] [CrossRef]
  34. Walsh, M.K.; Bombyk, R.A.; Wagh, A.; Bingham, A.; Berreau, L.M. Synthesis of lactose monolaurate as influenced by various lipases and solvents. J. Mol. Catal. B Enzym. 2009, 60, 171–177. [Google Scholar] [CrossRef]
  35. Chen, Y.; Nummer, B.; Walsh, M.K. Antilisterial activity of lactose monolaurate in milk, drinkable yogurt and cottage cheese. Lett. Appl. Microbiol. 2014, 58, 156–162. [Google Scholar] [CrossRef] [PubMed]
  36. Kim, J.E.; Han, J.J.; Yoon, J.H.; Rhee, J.S. Effect of salt hydrate pair on lipase-catalyzed regioselective monoacylation of sucrose. Biotechnol. Bioeng. 1998, 57, 121–125. [Google Scholar] [CrossRef]
  37. Ferrer, M.; Cruces, M.A.; Bernabe, M.; Ballesteros, A.; Plou, F.J. Lipase-catalyzed regioselective acylation of sucrose in two-solvent mixtures. Biotechnol. Bioeng. 1999, 65, 10–16. [Google Scholar] [CrossRef]
  38. Casas-Godoy, L.; Duquesne, S.; Bordes, F.; Sandoval, G.; Marty, A. Lipases: An overview. In Lipases and Phospholipases: Methods and Protocols; Sandoval, G., Ed.; Humana Press: Totowa, NJ, USA, 2012; pp. 3–30. [Google Scholar]
  39. Pleiss, J.; Fischer, M.; Schmid, R.D. Anatomy of lipase binding sites: The scissile fatty acid binding site. Chem. Phys. Lipids 1998, 93, 67–80. [Google Scholar] [CrossRef]
  40. Reyes-Duarte, D.; López-Cortés, N.; Ferrer, M.; Plou, F.J.; Ballesteros, A. Parameters affecting productivity in the lipase-catalysed synthesis of sucrose palmitate. Biocatal. Biotransformation 2005, 23. [Google Scholar] [CrossRef]
  41. Neta, N.S.; Teixeira, J.A.; Rodrigues, L.R. Sugar ester surfactants: Enzymatic synthesis and applications in food industry. Crit. Rev. Food Sci. Nutr. 2015, 55, 595–610. [Google Scholar] [CrossRef] [PubMed]
  42. Gumel, A.M.; Annuar, M.S.M.; Heidelberg, T.; Chisti, Y. Lipase mediated synthesis of sugar fatty acid esters. Process Biochem. 2011, 46, 2079–2090. [Google Scholar] [CrossRef]
  43. ICI Americas. The Hlb System: A Time-saving Guide to Emulsifier Selection; ICI Americas, Incorporated: New Castle, DE, USA, 1984; 19p. [Google Scholar]
  44. Hydrophile-lipophile balance of surfactants. Stud. Interface Sci. 2000, 9, 146–266. Available online: http://dx.doi.org/10.1016/S1383-7303(00)80016-2 (accessed on 3 September 2017).
  45. Zheng, Y.; Zheng, M.; Ma, Z.; Xin, B.; Guo, R.; Xu, X. Sugar fatty acid esters. In Polar Lipids; Elsevier: Amsterdam, The Netherlands, 2015; pp. 215–243. [Google Scholar]
  46. Naik, S.; Basu, A.; Saikia, R.; Madan, B.; Paul, P.; Chaterjee, R.; Brask, J.; Svendsen, A. Lipases for use in industrial biocatalysis: Specificity of selected structural groups of lipases. J. Mol. Catal. B Enzym. 2010, 65, 18–23. [Google Scholar] [CrossRef]
  47. Gennaro, A.R. Remington Farmacia; Editorial Médica Panamericana: Mexico City, Mexico, 2003; Volume 2, p. 2506. [Google Scholar]
  48. Griffin, W.C. Calculation of HLB values of non-ionic surfactants. Am. Perfum. Essent Oil Rev. 1954, 65, 26–29. [Google Scholar]
  49. Griffin, W.C. Classification of surface-active agents by "HLB". J. Soc. Cosmet. Chem. 1949, 1, 311–326. [Google Scholar]
  50. Davies, J. A quantitative kinetic theory of emulsion type, i. Physical chemistry of the emulsifying agent, Gas/Liquid and Liquid/Liquid Interface. In Proceedings of the International Congress of Surface Activity, London, UK, 1957; pp. 426–438. [Google Scholar]
Figure 1. Acylation positions of several sugars with different biocatalysts.
Figure 1. Acylation positions of several sugars with different biocatalysts.
Catalysts 07 00263 g001
Figure 2. Studied enzymatic reactions. R1, R2, and R3 are the acylation positions in lactulose, where each R is a lauryl residue. When the synthesis of monoesters occurs, the acylation positions are 1-O-, 6-O-, or 6′-O-, diester: (1-O-, 6-O-), (1-O-, 6′-O-), or (6-O-, 6′-O-), and triester: (1-O-, 6-O-, 6′-O-).
Figure 2. Studied enzymatic reactions. R1, R2, and R3 are the acylation positions in lactulose, where each R is a lauryl residue. When the synthesis of monoesters occurs, the acylation positions are 1-O-, 6-O-, or 6′-O-, diester: (1-O-, 6-O-), (1-O-, 6′-O-), or (6-O-, 6′-O-), and triester: (1-O-, 6-O-, 6′-O-).
Catalysts 07 00263 g002
Figure 3. Product profile of the enzyme screening developed in thin layer chromatography (TLC) plates. Reaction conditions: biocatalyst (25 mg/mL), lactulose (30 mM), vinyl laurate (150 mM), T = 60 °C, t = 24 h, solvent: 20% DMF in 2-methyl-2-butanol (2M–2B). (a) Lines: (L) Lactulose, (1) Novozym 435, (2) Lipozyme TL IM, (3) Lipozyme from Mucor miehei 20% DMF, (4) Lipozyme from Mucor miehei 0% DMF, (5) Lipase from Rhizopus orizae (Lewatit 160), (6) Lipase from Rhizopus oryzae (LewROL-PEI), (7) Lipase Amano PS from Burkholderia cepacia, (8) Lipase from Rhizomucor miehei (Palatase® 20,000 L), (9) Lipase from porcine pancreas, (10) Protease from Bacillus licheniformis. (b) Lines: (1) Novozym 435, (2) Lipozyme TL IM, (11) Alcaligenes sp. lipase (QLG).
Figure 3. Product profile of the enzyme screening developed in thin layer chromatography (TLC) plates. Reaction conditions: biocatalyst (25 mg/mL), lactulose (30 mM), vinyl laurate (150 mM), T = 60 °C, t = 24 h, solvent: 20% DMF in 2-methyl-2-butanol (2M–2B). (a) Lines: (L) Lactulose, (1) Novozym 435, (2) Lipozyme TL IM, (3) Lipozyme from Mucor miehei 20% DMF, (4) Lipozyme from Mucor miehei 0% DMF, (5) Lipase from Rhizopus orizae (Lewatit 160), (6) Lipase from Rhizopus oryzae (LewROL-PEI), (7) Lipase Amano PS from Burkholderia cepacia, (8) Lipase from Rhizomucor miehei (Palatase® 20,000 L), (9) Lipase from porcine pancreas, (10) Protease from Bacillus licheniformis. (b) Lines: (1) Novozym 435, (2) Lipozyme TL IM, (11) Alcaligenes sp. lipase (QLG).
Catalysts 07 00263 g003
Figure 4. TLC analysis for products profile in a one-hour reaction, using vinyl laurate as acyl donor (150 mM) and lactulose (30 mM) in organic solvent. Lines: (1) Novozym 435, (2) Lipozyme TL IM, and (3) Lipozyme from Mucor miehei.
Figure 4. TLC analysis for products profile in a one-hour reaction, using vinyl laurate as acyl donor (150 mM) and lactulose (30 mM) in organic solvent. Lines: (1) Novozym 435, (2) Lipozyme TL IM, and (3) Lipozyme from Mucor miehei.
Catalysts 07 00263 g004
Figure 5. ESI graphs of the complete reaction of: (a) Novozym 435; (b) Lipozyme TL IM.
Figure 5. ESI graphs of the complete reaction of: (a) Novozym 435; (b) Lipozyme TL IM.
Catalysts 07 00263 g005
Figure 6. MS-ESI (+) spectra from the purified reaction products: (a) Monoester fraction 1 of Novozym 435; (b) Monoester fraction 2 of Lipozyme TL IM.
Figure 6. MS-ESI (+) spectra from the purified reaction products: (a) Monoester fraction 1 of Novozym 435; (b) Monoester fraction 2 of Lipozyme TL IM.
Catalysts 07 00263 g006
Figure 7. Structural conformation of the lactulose, evidencing the less steric-hindered C-1 and C-6′ C-OH positions.
Figure 7. Structural conformation of the lactulose, evidencing the less steric-hindered C-1 and C-6′ C-OH positions.
Catalysts 07 00263 g007
Figure 8. Time-course of the monoester synthesis of lactulose by selected enzymes and effect of temperature in monoester production. Reaction conditions: 30 mM lactulose and 150 mM vinyl laurate, 25 mg/mL of biocatalyst, 2M2B (20% DMF). ―◆―Novozym 435, 30 °C and --◇-- 60 °C; ―▲― Lipozyme from Mucor miehei, 30 °C and - -△- - 60 °C, ―■― Lipozyme TL IM, 30 °C and - -☐- - 60 °C.
Figure 8. Time-course of the monoester synthesis of lactulose by selected enzymes and effect of temperature in monoester production. Reaction conditions: 30 mM lactulose and 150 mM vinyl laurate, 25 mg/mL of biocatalyst, 2M2B (20% DMF). ―◆―Novozym 435, 30 °C and --◇-- 60 °C; ―▲― Lipozyme from Mucor miehei, 30 °C and - -△- - 60 °C, ―■― Lipozyme TL IM, 30 °C and - -☐- - 60 °C.
Catalysts 07 00263 g008
Figure 9. Effect of sugar/acyl donor molar ratio on lactulose monoester enzymatic synthesis in 2M–2B with 20% DMF. Reaction conditions: lactulose (30 mM) and vinyl laurate (30 or 150 mM), 25 mg/mL of biocatalyst. ―◆―Novozym 435, 1:1 and --◇-- 1:5; ―▲― Lipozyme from Mucor miehei, 1:1 and - -△- - 1:5, ―■― Lipozyme TL IM, 1:1 and - -☐- - 1:5.
Figure 9. Effect of sugar/acyl donor molar ratio on lactulose monoester enzymatic synthesis in 2M–2B with 20% DMF. Reaction conditions: lactulose (30 mM) and vinyl laurate (30 or 150 mM), 25 mg/mL of biocatalyst. ―◆―Novozym 435, 1:1 and --◇-- 1:5; ―▲― Lipozyme from Mucor miehei, 1:1 and - -△- - 1:5, ―■― Lipozyme TL IM, 1:1 and - -☐- - 1:5.
Catalysts 07 00263 g009
Figure 10. Lactulose acylation positions with Novozym 435 and Lipozyme TL IM.
Figure 10. Lactulose acylation positions with Novozym 435 and Lipozyme TL IM.
Catalysts 07 00263 g010
Table 1. Comparative NMR data of the experimental and predicted data of monoesters 1, 6, and 6′.
Table 1. Comparative NMR data of the experimental and predicted data of monoesters 1, 6, and 6′.
Catalysts 07 00263 i001
Monoester Fraction Monoester-1 (1-O) (ppm)Monoester-6 (6-O) (ppm)Monoester-6′ (6′-O) (ppm)
Atom13Cexp13Ccal1Hexp1Hcal13Cexp13Ccal1Hexp1Hcal13Cexp13Ccal
Fructose moiety164.0064.044.394.46 4.2163.5363.873.94 (dd)3.92 (dd)64.2164.42
2103.11103.01 104.62104.43 104.12104.43
487.0087.183.61 (dd)3.65 (dd)87.0187.183.60 (dd)3.63 (dd)86.988.20
663.7963.814.00, 3.583.99, 3.5863.8063.814.45, 4.32, 4.07--------
Galactose moiety1′103.97103.923.924.03103.96103.923.924.033.924.03
5′76.0876.69 74.9674.94
6′61.4361.723.70, 3.463.70, 3.5062.6362.564.14 (dd)4.13 (dd)4.424.32 4.07
Lauroyl moiety1″173.5173.5 174.1174.1 173.8173.5
2″33.5834.032.38 (m)2.34 (m)33.5634.032.392.37 (m)
13Cexp: Experimental data in ppm; 13Ccal: Predicted data in ppm. 1Hexp: Experimental data in ppm; 1Hcal: Predicted data in ppm. Prediction calculated with: MestReNova Version 6.0.2–5475 and ACD/NMR Predictor Version 9.02 (ACD-Labs, USA).
Catalysts EISSN 2073-4344 Published by MDPI AG, Basel, Switzerland RSS E-Mail Table of Contents Alert
Back to Top