JOURNAL OF BIOENGINEERING AND RESEARCHcmibq.org.mx/jbbr/images/docs/jbbr-vol-3-no-2.pdfBautista Ramírez, Germán F. Gutiérrez Hernández, Elizabeth Argüello García, Martínez Herrera

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JOURNAL OF BIOENGINEERING AND BIOMEDICINE RESEARCH, Año 3, Volumen 3, No. 2, mayo - agosto 2019, es una Publicación cuatrimestral editada por el Colegio Mexicano de Ingenieros Bioquímicos, A.C.,Calle Mar del Norte #5, Col. San Álvaro, Alcaldía Azcapotzalco, Ciudad de México, C.P. 02090, Tel. (55)2873 2956, www.cmibq.org.mx, [email protected], [email protected] Editor Responsable: Deilia Ahuatzi Chacón, Rosa María Ribas Aparicio.Reserva de derechos al uso exclusivo No. 04-2016-041313084800-203, ISSN: 2594-052X, ambos otorgados por el Instituto Nacional del Derecho de Autor. Responsable de la última actualización de este Número, José Alberto Romero León Prolongación de Carpio y Plan de Ayala s/n, Col. Santo Tomás,Alcaldía Miguel Hidalgo, C.P. 11340, Ciudad de México fecha de última modicación 31 de mayo de 2019




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INDEX

area authors pages

Food technology

Bioactive Natural Products

1-7

8-16

17-27

Corzo Rios Luis Jorge, Esther Bautista Ramírez, Germán F. Gutiérrez Hernández, Elizabeth Argüello García, Martínez Herrera Jorge

Ruiz Ruiz Jorge Carlos, Us Medina Ulil, Segura Campos Maira Rubí

Herrera Pool Emanuel, Patrón Vázquez Jesús, Ramos Díaz Ana, Ayora Talavera Teresa, Pacheco López Neith

Technofunctional properties of the our and protein concentrate of canola (Brassica napus)

Combined process of liquid solid extraction, occulation,and adsorption to obtain an extract with high content of steviol glucosides

Extraction and identication of phenolic compounds in roots and leaves of Capsicum chinense by UPLC–PDA/MS

Food Science

28-39Parada Rabell Francisco, Chavez Parga Maria del Carmen, Alvarez B. Valente

Chemical stability and residual plasmin activity of milk subjected to various pressure-temperature treatments

Copyright ©2019 COLEGIO MEXICANO DE INGENIEROS BIOQUIMICOS

JOURNAL OF BIOENGINEERING AND BIOMEDICINE RESEARCH (2019) Vol.3 No.2 1-7

1

Research article

Technofunctional properties of the flour and protein concentrate of canola (Brassica napus)Corzo Rios Luis Jorge1, Esther Bautista Ramírez1, Germán F. Gutiérrez Hernández1, Elizabeth Argüello García2, Martínez Herrera Jorge *3

1 Unidad Profesional interdisciplinaria de Biotecnología, Dpto. de Bioprocesos. Instituto Politécnico Nacional. Av. Acueducto s/n, Col. Ticomán. Ciudad de México. C.P. 07340. México. 2 Universidad Popular de la Chontalpa. Cárdenas, Tabasco. México3 Instituto Nacional de Investigaciones Forestales, Agrícolas y Pecuarias, Campo Experimental Huimanguillo, Km. 1. Carr. Huimanguillo-Cárdenas, Tabasco, 86400, México

*Corresponding author: [email protected]

Received: 08 april 2019/ Accepted: 17 may 2019/ Published online: 31 may 2019

Abstract. Canola (Brassica napus) seeds are characterized by their high protein and oil content. Canola “press cake” after extraction of oil contains 36-44% protein. An alternative for the revaluation of these by-products is the application of modern technologies that add value through the transformation into products with characteristics and properties that allow their use in human nutrition, as they are isolated and protein concentrates. The objective in this work was to evaluate the techno-functional properties of canola (Brassica napus) flour and protein concentrate. From degreased canola flour (CF), the canola protein concentrate (CPC) was obtained by alkaline solubilization and isoelectric precipitation. In the CF the emulsifying capacity was higher at pH 2 (85.12%), in the case of CPC, it was higher at pH 2 (92.6%) and pH 8 (91%). The emulsifying stability in the CPC was strongly affected by the pH, but not in the CF. The foaming capacity in CPC was higher at pH 2 (298%) and pH 8 (277%), in CF it was at pH 2 (159%). The foam stability was higher in CPC than in CF. In general, the techno-functional properties evaluated had higher values in CPC than in CF.

Keywords. Canola, Foam, Emulsion.

Resumen. Las semillas de canola (Brassica napus) se caracterizan por su alto contenido de proteínas y aceite. La pasta de canola después de la extracción del aceite contiene entre 36 y 44% de proteína. Una alternativa para la revalorización de los subproductos es la aplicación de tecnologías modernas que agregan valor a través de la transformación en productos con características y propiedades que permita su uso en la alimentación humana. El objetivo del trabajo fue evaluar las propiedades tecno-funcionales de la harina de canola y concentrado de proteína. A partir de harina de canola desengrasada (HC), se obtuvo el concentrado de proteína de canola (CPC) por solubilización alcalina y precipitación isoeléctrica. En el CF, la capacidad emulsionante fue mayor a pH 2 (85.12%), el CPC fue más alto a pH 2 (92.6%) y pH 8 (91%). La estabilidad emulsionante en el CPC se vio afectada por el pH, pero no en la HC. La capacidad de espuma en CPC fue mayor a pH 2 (298%) y pH 8 (277%), en HC el mayor valor se obtuvo a pH 2 (159%). La estabilidad de la espuma fue mayor en CPC que en HC. En general, como era de esperarse, las propiedades tecno-funcionales presentaron valores más altos en CPC que en HC.

Palabras clave. Canola, Espuma, Emulsión.



2

INTRODUCTION

The canola was created in the late seventies, the rapeseed oil industry in Canada adopted this name to identify the cultivars of Brassica napus and Brassica campestris that, genetically, had a low content of erucic acid and glucosinolates.1

The canola (Brassica napus) is a winter oilseed and is part of the cruciferous family. Canola seed has 36 to 42% oil and is used for the production of edible and industrial oils, which is why it is a seed that is highly valued by industries. The by-product that is obtained after the extraction of the oil, is the “press cake”, which has an average content between 36 to 44% of protein and 12 a 13% of crude fiber.2,3

The functional properties of proteins are frequently affected by protein solubility; the most affected are the thickening, foaming, emulsifying and gelling properties. Insoluble proteins have a very limited use in foods. The solubility of a protein is the thermodynamic manifestation of the balance between protein-protein and protein-solvent interactions. The interactions that most significantly influence the solubility characteristics of proteins are hydrophobic and ionic.4

Hydrophobic interactions promote protein-protein association and decrease solubility, while ionic interactions promote protein-water interactions and increase solubility. The solubility of a protein is imposed by the hydrophilicity and hydrophobicity of the surface of the same that contacts the surrounding water and not with the average hydrophobicity and load frequency of the molecule as a whole. The smaller the number of hydrophobic zones on the surface, the greater the solubility. The solubility is also influenced by intrinsic physical-chemical properties, by

the pH, the ionic strength, the temperature, the presence of organic solvents and other concurrent circumstances in the solution.5

An alternative for the revaluation of these by-products is the application of modern technologies that add value through the transformation into products with characteristics and properties that allow their use in human nutrition, such as isolates and protein concentrates. They have appropriate technofunctional properties allowing their use in suitable amounts and enhancing the organoleptic characteristics of the food to which they are added. Therefore, the objective of this work was to evaluate the influence of pH and the purification process on the techno-functional properties of canola proteins (Brassica napus).

MATERIALS AND METHODS

Raw material

For the development of experimental activities, canola flour (CF), donated by the company Industrial Aceitera S.A. of C.V., was used as a source of protein. The flour was milled using a disk mill until the sample passed the screen 50 mesh (300 μm).

Proximate Composition

The proximate composition was determined according to the AOAC6 International methods, namely nitrogen (954.01); fat (920.39); ash (923.03); crude fiber (962.09); humidity (925.09); and Nitrogen free extract was calculated by differences.

Canola protein concentrate (CPC)

CPC was obtained according to the method described by Velazquez et al. 7 100 g of CF were dispersed in 1 L of distilled water,



3

The pH 12 was adjusted by 1N NaOH, left stirring constantly for 30 minutes, filtered on a screen, 100 mesh aperture (150 μm). After centrifugation at 3200Xg, for 10 minutes in a Centrifuge HERMLE Z260A, the sediment was discarded and the supernatant liquid was adjusted to pH 4 with 1N HCl, and the protein was recovered by centrifugation. The CPC was lyophilized at -47 ° C and 13x10-3 mbar.

Foaming Capacity

The foaming capacity was determined according to the method described by Corzo-Rios8. Suspensions containing 1% (w / v) of protein were adjusted to the pH values 2, 4, 6 and 8 (with 1.0 N NaOH or 1.0 N HCl). Then, they were stirred for 60 seconds at a temperature of 25° C, using a commercial blender, then transferred to a test tube to measure the volume.

The foaming capacity (FC) was determined according to the following equation:

(Total volume after shaking - foam volumen)

(Initial volume)

Foam stability

The method proposed by Corzo-Rios8 were used, following the same procedure used to determine the foaming capacity, with times of 30 seconds, 5, 15, 30, 60 and 120 minutes of rest at room temperature (25 ° C).

Emulsifying capacity

An adaptation of the technique recommended by Jiménez-Colmenero and García-Matamoros9 was used to determine the emulsifying activity. A 0.4% protein suspension was prepared. The pH was adjusted to different values (2, 4, 6, 8) with a 0.1N NaOH and 0.1N HCl solutions.

A commercial blender was used in which two electrodes were fixed, which were connected to a multimeter. The suspension was placed in the blender jar, the multimeter was connected and homogenized at low speed for 45 seconds. Later, it was brought to maximum speed and oil was added at a constant flow rate. The collapse of the emulsion was detected by an abrupt decrease in electrical resistance. At this time when the oil was stopped, the multimeter was disconnected, and the amount of oil used was recorded.

(mL of oil spent in the emulsion)

(Volume of sample dispersion)

Emulsifying stability

10 mL of oil were placed in 30 mL of the suspension at a concentration of 3 mg / mL and the pH was adjusted to 2, 4, 6 and 8 with solutions of 0.1 N NaOH and 0.1 N HCl. It was homogenized in a blender at high speed for 30 seconds. The emulsion was immediately poured into a 10 mL graduated cylinder. After the 10 mL in the test tube was completed, the time recording and the observation of the interface that appeared as result of the separation of a lower phase and a superior phase were initiated.

The kinetics of this process was quantified by measuring the volume of aqueous phase at 30 seconds, 5, 15, 30 and 120 minutes.

All determinations of foaming and emulsifying capacity and stability were made in triplicate and the values of the means are reported.

Statistical analysis.

Significant differences between the means were estimated using Duncan’s multiple range tests considered significant at p ≤ 0.05.

FC= x100

x100EC=



4

RESULTS

Chemical composition of flour and protein concentrate. As shown in Table 1, residual canola flour presented 38.22% protein, similar to that reported by Velázquez et al.7 and Aguilera Barreyro et al.,10 (38.2 and 38.1%) the content of fat, as expected, is in low percentage (2.75%) because canola flour was used as a by-product of the industrial process of oil extraction. On the other hand, a content of 79.22% of protein was achieved in the canola protein concentrate. It is also observed that in the canola protein concentrate it was possible to reduce the content of fiber and NFE in 60 and 78% respectively, until obtaining contents of 5 and 8.58%.

Table 1: Proximate composition of Canola Flour (CF) and Canola Protein Concentrate (CPC) (g/100g of dry matter)Component CF CPCMoisture 5.26 ± 0.30 4.50 ± 0.20

Protein 38.22% ± 0.05 79.22 ± 0.95

Fat 2.75% ± 0.14 1.50 ± 0.12

Fiber 12.17% ± 1.43 5.00 ± 0.30

Ash 1.76% ± 0.52 1.20 ± 0.10

NFE* 39.84 ± 2.95 8.58 ± 0.55Data are mean values of triplicate samples ± SD. *Nitrogen Free Extract (NFE) was calculated by difference

Figure 1 shows the results obtained at different pH values for the foaming capacity of canola flour and CPC, demonstrating that as the pH values is near to the isoelectric region it decreases its capacity to foam. Velazquez et al. 7 reported the isoelectric point of canola proteins at pH 4.

It is also observed that the highest foaming capacity was obtained for CPC at acidic pH values (pH 2) and at alkaline pH values (pH 8), with statistically equal foaming capacity values (p> 0.05). While, for canola meal, the foaming capacity was higher at alkaline pH and at other pH values this capacity was statistically the same.

Fig. 1: Foaming capacity from CF and the CPC at different pH values.

The foam stability of the CPC is observed in Figure 2, the greatest instability occurs at pH 2, greater reduction from 30 seconds to 120 minutes. It may be due to the fact that the protein film formed around each gas bubble was not sufficiently viscous, elastic and resistant to reduce the permeability of the gas, inhibit the coalescence of the foam, and therefore incapable of supporting the weight of the foam formed, since at that pH the highest volume of foam was obtained.5, 11

Considering all the results, both the foaming capacity and the stability of the foam are increased as they approach alkaline pH values (figure 1 and 2).

Fig. 2. Foam stability in the CPC at different pH with respect to time.

In the same way, the emulsifying properties of the CPC are strongly affected by the pH values. The highest values of emulsifying capacity were obtained at pH values 2 and 8 (91 and 90% respectively), and at pH 4 the emulsifying capacity was low (55%). (Figure 3).

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In addition, it was found that the canola f lour (CF) emulsifying capacity was higher at pH 2 and lower at pH 6 and 8, with the foaming capacity at pH closer to IP of canola f lour being greater than that presented by the protein concentrate.

In figure 4, it is observed that the emulsifying stability of the CPC is very higher, in the values of pH 2 and 8 with emulsifying capacities of 100% and 90% respectively. On the other hand, at the pH values near the isoelectric point, the emulsifying stability was low.

Fig. 4. Emulsifying stability in different pH, with respect to the time from CPC.

DISCUSSION

The protein content in CPC indicates its potential use as source of ingredients for human food, in addition, the low fat and NFE content suggests a low caloric intake. The method used to obtain the protein concentrate is simple and a higher protein concentration than that reported by other researchers for protein legume concentrates was achieved.12

As in the emulsion, the ability of the protein to develop foam depends on its active surface properties, although to a different amount.11 In this way, as the concentration of protein (from CF to CPC) increases, the foaming capacity also increases.

This can be explained due to the denaturation that the proteins suffer during the process of obtaining the CPC, which would expose regions of hydrophilic amino acids which would increase the ability to stabilize the air-water interface. During the formation of the foam, the functional protein is concentrated in the gas bubble and the liquid interface reducing the surface tension thus increasing the viscosity. This keeps the gas bubble and minimizes the waste of the liquid, but to have a satisfactory performance part of the protein must be initially soluble and able to unfold from the interface.5,13

The good foaming capacity at acid pH values suggests the possibility of using the canola protein concentrate in products such as meringues and mousses. On the other hand, because of the good stability shown at basic pH values, its use it in baked goods such as breads, cakes and biscuits is suggested, since in the latter it would act as a support medium for the starch, achieving a maximum volume before baking.4

The lower percentages of emulsifying capacity were observed in the isoelectric region, but not in the canola flour where it has an elevated emulsifying capacity at the pI with 74%, still higher at pH 6 and 8 with a capacity of 62% and 65% respectively. This could be due to the interaction of the protein with carbohydrates present in the flour, which can interfere with the surface properties of the protein and the possibility of interacting the regions of hydrophobic amino acids with the oil phase.11, 13

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The low solubility of some proteins in the isoelectric region can decrease the emulsifying capacity, since these adopt a compact structure that prevents the splitting and absorption at the interface, which is not desirable in an emulsion. These emulsifying properties improve as they move away from the isoelectric region, which can be attributed to an increase in electrical charge that increases protein solubility.5,13

In the evaluation of protein concentrates of legumes, the emulsifying capacity values range from 53 to 58%.14 Pérez et al.15 reports emulsifying capacity values at different pH values for flours and protein concentrates of Phaseolus lunatus and Canavalia. ensiformis ranging from 40 to 60%. On the other hand, Cano-Medina et al.16 report an emulsifying capacity of 38 y 44% for the sesame and soy isolates, respectively; these values being lower than those obtained in the CPC with a maximum value of 93%.

The results of emulsifying stability shown in the figure 4, clearly indicate that the canola meal and of the CPC is greater stability, in the CPC this due to the availability of the proteins in this same stability is much higher, from 30 seconds to 120 minutes, at pH values 2 and 8 with an emulsifying capacity of 100% and 90% respectively.

Chau et al.14 obtained, for legume protein concentrates, emulsion stability values of 94.8% at pH 4, this being the lowest value reported and values of 96 and 97% at pH 2 and 10, respectively. Pérez et al. 15 reported 100% emulsifying stability values for flours and protein concentrates of P. lunatus and C. ensiformis. The stabilization of the emulsion is achieved by restricting the mobility of the droplets of the dispersed phase, thanks to the increase in viscosity and sometimes the viscoelasticity of the continuous phase.5,16 It has been observed that the behavior of the elasticity of the protein layers adsorbed at the oil-water interface can affect the stability of an emulsion.13,16

Different authors report values of foaming and emulsifying capacity that coincide with the results obtained for canola proteins. The best performance of these technofunctional properties are obtained at pH values far from the isoelectric point, which is totally congruent; because these properties are related to ability to diminish the interfacial tension, existing a greater disposition of the molecules to act in the interface.5,11,13

CONCLUSIONS

The foaming and emulsifying properties of canola proteins increased as the purity was higher, that is, the functional properties were better in CPC than in CF, and pH had a greater effect on these properties in CPC than in CF, as an additional consequence of the purification of proteins.

FUNDING AND ACKNOWLEDGMENTS

This research was supported by SIP-IPN through project support with no. 20140215. Thanks are due to Instituto Politécnico Nacional, COFAA-IPN and EDI-IPN.

REFERENCES

1) Romero A.H. 1993. Evaluación de rendimiento y características agronómica de la colza y-o canola (Brassica spp) en Tepatitlán Jalisco. Tesis de licenciatura, Universidad de Guadalajara, Guadalajara, México.

2) Canola Council. 2009. Pasta de canola. Guía para la industria de forrajes.: http://www.canolacouncil.org /media/516719/canola_meal_feed_guide.pdf. 09-04-15.

3) Gonçalves N, Vioque J, Clemente A, Sánchez-Vioque R, Bautista J. Millán F. 1997. Obtención y caracterización de aislados proteicos de colza. En: Grasas y Aceites, Vol. 48: 282-289.

4) Damodaran S. 2005. Food Proteins and Their



7

Applications. J. Food Sci 70(3): 54-66.5) Damodaran, S. and Song K. B. 1988. Kinetics

of adsorption of proteins at interfaces: role of protein conformation in diffusional adsorption. Biochimica et Biophysica Acta (BBA) - Protein Structure and Molecular Enzymology. 954: 253-264.

6) AOAC, 1984. Official Methods of Analysis. Association of Official Analytical Chemists. 13ª. Ed. Washington.

7) Velázquez-Sánchez I, Jiménez-Martínez C, Dávila- Ortiz G, Corzo-Rios L. 2016. Evaluación de los cambios en algunas propiedades bioactivas de proteínas de canola (Brassica napus) con la hidrólisis enzimática. XX Congreso Nacional de Ingeniería Bioquímica. Veracruz, México.

8) Corzo-Rios L.J. 2014. Evaluación de las propiedades de superficie y mecánicas de sistemas mixtos de goma carboximetilada de flamboyán (Delonix regia) con hidrolizados proteicos de Phaseolus lunatus L. Tesis Doctorado FIQ-UADY.

9) Jiménez-Colmenero F, and García-Matamoros E. 1981. Effects of washing on the properties of mechanically deboned meat. In proceedings of the 27th European meeting of meat research workers. O. Pränld (Ed.) Vienna. pp 351-354.

10) Aguilera-Barreyro A, Reis de Souza T, García GT, Gómez SJ, Escobar GK, Bernal-Santos M, Montaño BS. 2011. Composición química, constituyentes antinutricionales y fracciones de proteína de pastas de ajonjolí, soya y canola. XV Congreso Bienal AMENA.

11) Betancur-Ancona D, Gallegos-Tintoré S, Chel-Guerrero L. 2004. Wet-fractionation of Phaseolus lunatus seeds: partial characterization of starch and protein. J Food Sci Agric. 84 (10): 1193–1201.

12) Wagner J. 2000. Propiedades superficiales en Caracterización funcional y estructural de proteínas. Editores, Pilosof, A y Bartholomai

G. Editorial, Universidad de Buenos Aires.13) Damodaran S. 2005. Protein Stabilization of

Emulsions and Foams. J. Food Sci. 70(3): 54-66.

14) Chau C, Cheung P, Wong Y. S. 1997. Functional properties of protein Concentrates from fhree chinese indigenous legume seeds. J Agr Food Chem. 45(7):2500-2503.

15) Pérez, F. V., Chel, G.L., Betancur, A.D. and Dávila,O.G. 2002. Functional properties of flours and protein concentrates from Phaseolus lunatus and Canavalia ensiformis seeds. J Agr Food Chem. 50(3): 584–59.

16) Cano-Medina A, Jiménez-Islas H, Dendooven L, Patiño-Herrera R., González-Alatorre G., Escamilla-Silva E. M. 2011. Emulsifying and foaming capacity and emulsion and foam stability of sesame protein concentrates. Food Res Int. 44 (83):684-692.

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Research article

Combined process of liquid solid extraction, flocculation, and adsorption to obtain an extract with high content of steviol glucosides Ruiz Ruiz Jorge Carlos1, Us Medina Ulil2, Segura Campos Maira Rubí2*

1Escuela de Nutrición, División de Ciencias de la Salud, Universidad Anáhuac-Mayab. Km 15.5 Carretera Mérida a Progreso, Int. Km 2 Carretera a Chablekal. Mérida, Yucatán, México. C.P. 97310. Tel. +52 (999) 942-48-00. 2Facultad de Ingeniería Química, Campus de Ciencias Exactas e Ingenierías, Universidad Autónoma de Yucatán. Periférico Nte. Km. 33.5, Tablaje Catastral 13615, Col. Chuburná de Hidalgo Inn, 97203. Mérida, Yucatán, México. Tels. +52 (999) 9460956, 946-09-81 and 946-09-89; Fax. +52 (999) 946-09-94.

Corresponding author: [email protected]: 08 january 2019/ Accepted: 13 march 2019/ Published online: 31 may 2019

Abstract. A process of solid-liquid extraction, flocculation and adsorption was developed to obtain an extract with a high content of steviol glycosides, from Stevia rebaudiana, leaves. The process started with an aqueous extraction of the of the ground leaves. Different solid-liquid relationships, temperatures and extraction times were evaluated. Then a precipitation with calcium chloride and a stage of adsorption with celite were carried out to remove components that generate color and turbidity. Considering the response variables, steviol glycoside content, color, turbidity and soluble solids content, it was established that the best conditions for extraction were a solid-liquid ratio (1:5) during 30 min at 90 °C. For flocculation, the best results were obtained using calcium chloride (10 mg/mL) for 30 min at pH 2. The best results for adsorption were obtained using celite (1 mg/mL) for 10 min. After the three-stages process, the concentration of glycosides increased by 21%. The extract was crystallized under reduced pressure at 50 °C, obtaining a concentration of glycosides of 284.2 mg/g, consisting mainly by Rebaudioside A (98.5%).

Keywords: Steviol glycosides, Stevia rebaudiana,, solid-liquid extraction, flocculation, adsorption.

Resumen. Se desarrolló un proceso de extracción sólido-líquido, floculación y adsorción para la obtención de un extracto con alto contenido de glucósidos de esteviol, a partir de hojas de Stevia rebaudiana,. El proceso inició con la extracción acuosa de las hojas molidas. Se evaluaron diferentes relaciones sólido-líquido, temperaturas y tiempos de extracción. Después se efectuó una precipitación con cloruro de calcio y una etapa de adsorción con celita para remover componentes que generan color y turbidez. Considerando las variables de respuesta, contenido de glicósidos de esteviol, color, turbidez y contenido de sólidos solubles, se estableció que las mejores condiciones para la extracción fueron una relación sólido-líquido (1:5) durante 30 min a 90 °C. Para la floculación, los mejores resultados se obtuvieron usando cloruro de calcio (10 mg/mL) durante 30 min a pH 2. Lo mejores resultados para la adsorción se obtuvieron usando celite (1 mg/mL) durante 10 min. Después de las tres etapas del proceso, la concentración de glicósidos se incrementó un 21%. El extracto se cristalizó a presión reducida a 50 ºC, obteniéndose una concentración de glicósidos de 284.2 mg/g, constituida principalmente por Rebaudiosido A (98.5%).

Palabras clave: Glicósidos de Steviol, Stevia rebaudiana,, extracción sólido-líquido, floculación, adsorción.



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INTRODUCTION

Stevia rebaudiana, leaves contain steviol glycosides that human organism does not have the capacity to metabolize, for this reason does not generate an impact in glycemic index. Several studies indicate that the stevia consume not only has beneficial effects on the nutritional status of patients with diabetes, but also has antioxidant and hypotensive effects1. Therefore, steviol glycosides can be considered as high value-added natural compounds. In order to obtain this kind of compounds, five stages universal recovery process are employed: macroscopic pre-treatment, macro and micro molecules separation, extraction, isolation and purification, product formation.2,3

Processing technologies and specific methodologies have been developed for nutraceuticals purification. Wet milling, mechanical pressing and microfiltration for macroscopic pretreatment. Solvent precipitation, isoelectric solubilization and ultrafiltration for macromolecules remotion. Solvent and supercritical fluid extraction separate smaller molecules. Resin adsorption, chromatography and nanofiltration isolate and purify compounds before obtaining a final stabilized product.2

Several studies and patents have been published on processes of extraction and purification of steviol glycosides. The most used extraction processes are based on the use of solvents, solvents plus a bleaching agent, and supercritical fluids. As for purification, adsorption, chromatographic techniques and the use of membranes are the most used methods.4 These extractive and purification processes can generate residues harmful to health. In addition to being costly methodologies in terms of reagents and energy consumption.5,6

The present study is aimed on obtain an extract rich in steviol glycosides from the leaves of Stevia rebaudiana, by removing as much as

possible impurities, color and turbidity. A recovery process is proposed that combines the use of different methodologies, such as solid-liquid extraction, flocculation and adsorption.


Raw material and chemicals

Stevia rebaudiana, Bertoni variety Morita II was obtained from plots in Mocochá, Yucatan, Mexico. Samples were obtained from the first cut of the plant at three months age. Stevia rebaudiana, leaves were subjected to convection drying at 60 °C for 24 h. The leaves were milled to obtain particles of 1.0 mm in size. Samples were stored at room temperature in polyethylene bags until extract preparation. All chemicals were reagent grade or better and purchased from Sigma Chemical Co. (St. Louis, MO, USA).

Solid-liquid extraction experiments.

A 32 factorial model with four central treatments was used to evaluate the solid-liquid extraction conditions. The evaluated factors and levels were solid-liquid ratio (1:5 and 1:20 w/v), temperature (30 and 90 °C), and extraction time (30 and 90 min). Response variables included soluble solids content (SSC), color (C), turbidity (T) and steviol glycosides content (SGC). The extracts were prepared by mixing the stevia leaves powder with water (10 g/50 mL and 2.5 g/50 mL). Dispersion was heated (30 and 90 °C) and stirred (30 and 90 min) in a water bath shaker (MaxQ 700, Lab-Line, USA). After cooling, extracts were filtered through a paper filter and centrifuged in Hermle Z300K (Hermle Labortechnik, Wehigen, Germany) at 3000 rpm for 15 min. Finally, the extracts were stored in amber bottles and refrigerated at 4 °C until analysis. The extract with higher SGC and lower SSC, C, and T was selected to perform the chemical flocculation experiments.



Chemical flocculation experiments

A 32 factorial model with four central treatments was used to evaluate the chemical flocculation conditions. The evaluated factors and levels were flocculant agent concentration (10 and 50 mg/mL), pH (2 and 10), and time (10 and 30 min). Response variables included content of steviol glycosides, color, turbidity and content of solids. At a volume of 30 mL of extract was added the flocculant agent (10 and 50 mg/mL), pH was adjusted with NaOH 1.0 M or HCl 1.0 M and stirred (10 and 30 min). After, extracts were centrifuged in Hermle Z300K (Hermle Labortechnik, Wehigen, Germany) at 5000 rpm for 30 min to remove the flocculants. Finally, the extracts were stored in amber bottles and refrigerated at 4 °C until analysis. The extract with higher SGC and lower SSC, C, and T was selected to perform the adsorption experiments.

Adsorption experiments

A 22 factorial model with four central treatments was used to evaluate the adsorption conditions. The evaluated factors and levels were adsorbent agent concentration (1 and 10 mg/mL) and time (10 and 30 min). Response variables included content of steviol glycosides, color, turbidity and content of solids. At a volume of 20 mL of extract was added the adsorbent agent and stirred with a magnetic stirrer. After, extracts were centrifuged in Hermle Z300K (Hermle Labortechnik, Wehigen, Germany) at 5000 rpm for 30 min. Finally, the extracts were stored in amber bottles and refrigerated at 4 °C until analysis.

Soluble solids, color and turbidity determinationThe refractometric method established by Mexican regulations was used for determinate soluble solids.7 Optical absorbance was measured an evolution-220 spectrophotometer (Thermo Scientific, USA) at wavelengths of

420 and 670 nm was correlated with the color and turbidity removal parameters, respectively.8 These parameters were determined in triplicated for each sample after extraction, flocculation and adsorption experiments.

Quantification of steviol glycosides

Was performed using the DNS (3,5-dinitrosalicylic acid) method, that allows correlate the content of steviol glycosides with the carbohydrate content.9 A calibration curve was obtained with standard glucose concentrations in a range of concentrations from 0.0 to 1.2 mg/mL. The glucose content was measured in an evolution-220 spectrophotometer (Thermo Scientific, USA) at wavelength of 540 nm. This parameter was determined after extraction, flocculation and adsorption experiments in triplicated.

Quantification of individual steviol glycosides by HPLC

The quantification was carried out using the methodology proposed by the Joint FAO/WHO Expert Committee on Food Additives .10 To prepare the standard solutions, 50 mg of stevioside (Sigma S3572) and 50 mg of rebaudioside A (Sigma 01432) were weighed. Then were placed in 50 mL volumetric flasks and dissolved with a mixture of water-acetonitrile (7:3, v/v). To prepare the sample solutions, 100 mg of dry extract were accurately weighed. Then were placed in 50 mL volumetric flasks and dissolved with a mixture of water-acetonitrile (7:3, v/v). For chromatographic analysis 5 µL of standard or sample solution were injected in a column Luna (5μm C18, 100 A, Phenomenex) with a length of 250 mm; an inner diameter of 4.6 mm and a particle size of 5 mm. The column was coupled to a HPLC Agilent model 1200 with diode array detector (210 nm). The mobile phase consisted in a 32:68 (v/v) mixture of acetonitrile



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and 10 mmol/L of sodium phosphate buffer (pH 2.6) at a flow rate of 1.0 mL/min during 30 min. The peaks from the sample solution were identified by comparing the retention time with the peaks from the mixture of steviol glycosides standard solutions. The quantification was carried out in triplicated for each sample after extraction, flocculation and adsorption experiments.

Statistical analysis

For experiments of solid-liquid extraction and chemical flocculation were used a 32 factorial models with four central treatments, for adsorption experiment was used a 22 factorial model with four central treatments. All analyses were carried out in Statgraphics Centurion XV software (version 15.2.14).

RESULTS

Solid-liquid extraction experiments

Table 1. Soluble solids, color, turbidity and steviol glycosides content after solid-liquid extraction.

CT = Central Treatment. Different superscript in the same column indicates significant statistical difference (p < 0.05).

Experiments were performed according to a 32 factorial model and the ANOVA results for modelled responses are reported in Table 1.

For soluble solids content, solid-liquid ratio (negative effect) and temperature (positive effect) were significant, treatment 6 (1:5 ratio, 30 min, and 90 °C) showed the higher content of soluble solids. Color was negatively affected by solid-liquid ratio and temperature, with treatment 1 (1:20 ratio, 30 min, and 30 °C) as the most colored. For turbidity, solid-liquid ratio (negative effect), time (positive effect), and temperature (positive effect) were significant, treatment 7 (1:20 ratio, 90 min, and 90 °C) showed the lower turbidity. Steviol glycosides content was negatively affected by solid-liquid ratio, with treatment 4 (1:5 ratio, 90 min, and 30 °C) as the highest concentration of steviol glycosides. Figure 1a shows the 3D surface plot of the interactive effects of the independent variables corresponding to the response of steviol glycosides content. It can be seen that the extraction yield of steviol glycosides of S. rebaudiana extracts increased with the decrease of solid-liquid ratio and the increase of the extraction time, reaching a maximum content of 1.0 mg/mL.


Table 2 shows the effect of the factors involved on chemical flocculation on the soluble solids, color, turbidity, and steviol glycosides percentage removal. The purpose of chemical flocculation is only to clarify the extract, so it is important to minimize the flocculation of steviol glycosides, thus the flocculant agent (CaCl2) must be able to flocculate substances that give color to the extract without flocculating stevia glycosides. Soluble solids were positively affected by flocculant concentration, with treatment 7 (50 mg/mL of flocculant, 30 min, and pH 10) as the

Treat-ment

So-lid-li-quid ratio

Time (min)

Tempera-ture (°C)

Soluble solids(ºBrix)

Color Turbi-dity

Steviol Glycosi-des (mg/

mL)

1 1:20 30 30 2.90a 0.08a 0.51a 0.56a

2 1:5 30 30 9.00bc 0.29b 1.67c 0.94b

3 1:20 90 30 2.00a 0.10a 0.59a 0.51a

4 1:5 90 30 9.00bc 0.34b 1.95c 1.00b

5 1:20 30 90 2.50a 0.10a 0.44a 0.60a

6 1:5 30 90 10.00c 0.18c 1.25b 0.94b

7 1:20 90 90 3.00a 0.09a 0.41a 0.58a

8 1:5 90 90 10.50c 0.20c 1.24b 0.88b

CT 1:10 60 60 7.00b 0.20c 1.33b 0.91b



treatment with lower content of soluble solids. For color, flocculant concentration and pH have a positive effect, central treatment (30 mg/mL of flocculant, 20 min, and pH 6) showed the higher reduction of color. Turbidity was negatively affected by pH, with treatment 1 (10 mg/mL of flocculant, 10 min, and pH 2) as the treatment with lower turbidity. For the content of steviol glycosides flocculant concentration (negative effect) and time (positive effect) were significant, treatment 3 (10 mg/mL of flocculant, 30 min, and pH 2) showed the higher content of steviol glycosides. Figure 1b shows the surface plot of the interactive effects of the independent variables corresponding to the response of steviol glycosides residuary content. It can be seen that the remotion of steviol glycosides of S. rebaudiana extracts increased with the increase of the flocculant agent and the flocculation time, reaching a maximum remotion of 34.69%.


Table 3 shows the effect of the factors involved on adsorption on the soluble solids, color, turbidity, and steviol glycosides percentage removal. Soluble solids were positively affected by time, with treatments 3 and 4 (10 mg/mL and 50 mg/mL of celite, and 30 min of adsorption) as treatments with higher content of soluble solids. For color, adsorbent agent concentration (positive effect) and time (negative effect) were significant, treatment 3 (1 mg/mL of adsorbent agent and 10 min) showed the higher reduction of color. Turbidity was positively affected by adsorbent agent concentration, with central treatment (5 mg/mL of adsorbent agent concentration and 15 min) as the treatment with lower turbidity. For the content of steviol glycosides adsorbent agent concentration (positive effect) was significant; treatment 1 (1 mg/mL of adsorbent

agent concentration and 10 min) showed the higher content of steviol glycosides.Figure 1c shows the surface plot of the interactive effects of the independent variables corresponding to the response of steviol glycosides residual content. It can be seen that the remotion of steviol glycosides of S. rebaudiana extracts increased with the increase of the adsorbent agent and time, reaching a maximum remotion of 49.52%.

Fig.1. 3D surface plot, interactive effects of the independent variables corresponding to the response of steviol glycosides content. a) Steviol glycoside content as function of solid-liquid ratio (w/v) and time (min): response surface. b) Steviol glycoside remotion as function of flocculant concentration (mg/mL) and time (min): response surface. c) Steviol glycoside remotion as function of adsorbent agent concentration (mg/mL) and time (min): response surface.

Higher concentration of adsorbent agent resulted in a reduction of the concentration of steviol glycosides.Considering the importance of the glycosides content, the contribution percentage of the



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variables was calculated. The solid-liquid ratio contributes 97% in the glycoside content response. Flocculant agent and flocculation time contributes 87% and 13%, respectively in the glycoside content response. The adsorbent agent concentration contributes 98% in the glycoside content response.

Table 2. Soluble solids, color, turbidity and steviol glycosides remotion after chemical flocculation.

CT = Central Treatment. Different superscript indicates significant statistical difference (p<0.05) in the same column.

Quantification of steviol glycosides by HPLCAfter all process the extract was saturated under reduced pressure at a temperature of 50 °C until the total crystallization. Crystals were ground to obtain a fine powder, placed in a crucible and their moisture removed in a desiccator. Finally, the steviol glycosides content was determined, to establish the purity and the yield of the process (Table 4).

After the first three stages of the process, concentration of steviol glycosides increased by 18.03%. The removal of some pigments and steviol glycosides from extract occurred during the formation of aggregates that subsequently precipitated. Results suggest that coagulation is the most relevant mechanism in the clarification of stevia extract.

Table 3. Soluble solids, color, turbidity and steviol glycosides remotion after adsorption.

CT = Central Treatment. Different superscript indicates significant statistical difference (p<0.05) in the same column.

Crystallized extract contains 280 mg/g of rebaudioside A and 4.2 mg/g of stevioside. This indicates that 28.42% of the solid material obtained after crystallization consists of steviol glycosides.

DISCUSSION

The results in present research contrast 11 Wong et al., whereby higher antioxidants compounds (phenolic and f lavonoids) content was obtained from palm kernel by-product, when liquid to solid ratio was increased. A higher liquid to solid ratio is preferred in extracting bioactive compounds from plant materials. This is because more solvent can enter cells and is helpful in permeating more compounds, which can diffuse in to the solvent under higher liquid to solid ratio.12

In this study, steviol glycosides content was positively correlated with the rest of the extract parameters (time and temperature); therefore, there is no single treatment suitable for obtaining the highest steviol glycosides content with low values of color and turbidity, simultaneously. The best process depends on the subsequent use as sweetener or an extract with less color and turbidity.

Treat-ment

CaCl2(mg/mL)

Time (min) pH

Soluble solids (ºBrix)

Color TurbiditySteviol

glycosides (%)

110 10 2

10.00c 55.53e -27.11a -13.88b

250 10 2

60.00g 4.85b -5.25c -34.69a

310 30 2

15.00d 9.16c 3.90d 15.60d

450 30 2

65.00h 72.24f -6.81b -17.90b

510 10 10

0.00b 185.44h 138.85f 7.95c

650 10 10

40.00e 176.01g 138.85f -18.67b

710 30 10

-5.00a 43.13d 138.85f 14.83cd

850 30 10

40.00e 236.66i 138.85f -12.73b

CT30 20 6

50.00f -25.61a 49.60e -10.66b

TreatmentAbsorbent

concentration(mg/mL)

Time (min)

Soluble solids Color Turbidity

Steviol glycosides

(%)

1 1.0 10 13.04a 90.77b 23.18a 5.05d

2 10.0 10 13.04a 134.32e 39.18c -49.52c

3 1.0 30 17.39b 67.51a 28.34b -40.54a

4 10.0 30 17.39b 116.27d 29.62b -48.22c

CT 5.0 20 13.04a 105.84c 21.40a -43.10b



Table 4. Content of steviol glycosides, rebaudioside A and stevioside during process.

Some authors13,14,15 refer the extraction method proposed by Kohda et al.16 which involved four main steps: extraction into polar organic solvent, discoloration, coagulation or concentration, column chromatography and crystallization. During the first step (extraction), powdered stevia leaves are soaked in petroleum ether followed by extraction with methanol for 3 to 4 times, then the methanol extract is concentrated under vacuum and the suspension is dissolved in distilled water. The extraction method proposed in present research involves an individual process which combines water in a low ratio (1:5), a moderate temperature (30 ºC) and time (90 min). In this way using treatment 4 is feasible to obtain a good yield of steviol glycosides, keeping to the minimum the consumption of reagents and energy, obtaining at the end the same result of conventional extraction method, a stevia aqueous extract.


In this research, the concentration of CaCl2 as flocculant as well as the flocculation time were significant for the studied response variables. In this sense, flocculants are chemicals that are used to promote flocculation by causing colloids and other suspended particles in liquids to aggregate, forming a floc. Many flocculants are multivalent cations; these positively charged molecules interact with negatively charged particles and

molecules to increase aggregation. In addition, many of these chemicals react with water to form insoluble hydroxides, which upon precipitating, link together to form meshes, physically trapping small particle into the larger floc.17 This would explain the behavior observed during the flocculation experiments of the extract of S. rebaudiana.

Cationic polyelectrolytes could be used for the whitening of stevia extracts, because positive charges can interact electrostatically with a counter-ion and then precipitate together with pigments. In this sense, Oliveira et al.8 tested positive charged chitosan (660 mg of chitosan/mL of stevia extract) and sodium tripolyphosphate (2 mL, 1% w/v) for flocculate pigments of stevia extracts. The results showed that chitosan was able to flocculate both the pigments and the steviol glycosides, with average removals of about 44.56% of glycosides. In present study CaCl2, proposed as flocculant agent, at a concentration of 10 mg/mL of stevia extract, only removes 15.06% of glycosides together with soluble solids (15.00%), color (10%) and turbidity (3.90%). Flocculant method proposed in this research only use one reactive (CaCl2) at low concentration (10 mg/mL) during 30 min and eliminates the use of sodium tripolyphosphate (ecotoxic). This could be considered as an advantage in terms of cost of reagents and the reduced environmental impact.


Adsorbent agent concentration was significant for color, turbidity and steviol glycosides content. Celite is an acid washed, high purity, flux calcined, due to its specific properties (porous structure, high silica content, low density, low conductivity coefficient, etc.); has extensively been applied in many ways, such as filter aid and adsorbent agent. Adsorption occurs when an impurity is physically taken onto the

Process Steviol glycosides content

Solid-liquid extraction

DNS method (mg/mL)

1.00

Chemical flocculation 1.16

Adsorption 1.22

Crystallization

HPLC quantification

Rebaudioside A (mg/g extract) 280

Stevioside (mg/g extract) 4.2



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surface of the filter aid particle. The impurity is than removed from the liquid system when the particle is filtered out of the solution.18 Among the methods employed for isolation and purification of bioactive compounds such chromatography, magnetic fishing, nanofiltration, membrane ion exchange chromatography,19 adsorption of bioactive compounds in filter particles could be is less selective and efficiency. Results indicated that experimental conditions for the use of celite as adsorbent agent material was not adequate as it significantly reduces the content of steviol glycosides in the extract. On the other hand, it does not have a positive effect on the reduction of color or turbidity.

Quantification of steviol glycosides by HPLC

The quantification of glycosides using HPLC chromatography indicated a significant increase of Rebaudioside A, until a ratio of Rebausioside A: Stevioside of 67:1 was reached. Usually the ratio of these two glycosides is 10:1.15 It is convenient to have a higher concentration of Rebaudioside A, since this compound is sweet and does not have the bitter taste of stevioside. The results indicated that process of solid-liquid extraction, flocculation and adsorption obtains mainly Rebaudioside A.

CONCLUSIONS

An extraction with a solid-liquid ratio of 1:5 (w/v) during 90 min at 30 °C allows obtaining an extract with low content of total solids, color and turbidity. At the same time, a high content of steviol glycosides is achieved. CaCl2 acted properly as chemical flocculant agent in the clarification of stevia extract. However, there is no single treatment suitable for obtaining the highest steviol glycosides content with low values of color and turbidity, simultaneously. The best treatment for chemical flocculation exhibited

low percentages of color (9.16%) and turbidity (3.90%) remaining. Adsorption negatively affected the content of steviol glycosides. Once the process is completed the concentration of steviol glycosides increased by 18.03%. Crystallized extract contains 28.42% of steviol glycosides, mainly Rebaudioside A (98.5%). The sequential use of aqueous extraction, chemical flocculation and adsorption as recovery process for steviol glycosides from Stevia rebaudiana, leaves, could be less expensive in terms of chemical reagents and technology requirements than other recovery methods such polar solvent extraction, flocculation with cationic polymers and chromatography. This is an important criterion for future studies focused on optimizing and making more efficient the process of recovery of steviol glycosides proposed in this research.

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Research article

Extraction and identification of phenolic compounds in roots and leaves of Capsicum chinense by UPLC–PDA/MS Herrera-Pool Emanuel1, Patrón-Vázquez Jesús1, Ramos-Díaz Ana1, Ayora-Talavera Teresa1, Pacheco Neith1*

1Centro de Investigación y Asistencia en Tecnología y Diseño del Estado de Jalisco A. C., Carretera Sierra Papacal-Chuburna Km 5, C. P. 97302, Sierra Papacal, Yucatán, México.


Received: 13 april 2018/ Accepted: 02 may 2019/ Published online: 31 may 2019

Abstract. Extractable phenolics (EPs) and non-extractable phenolic (NEPs) compounds were obtained by using ultrasound-assisted extraction (UAE) with different polarity solvents from roots and leaves of Capsicum chinense seedlings. Significant differences were not found in total phenolic compounds (TPC) from EPs. However, a higher TPC was obtained by using acid extraction rather than alkaline when working with NEPs. Furthermore, a higher EPs content was observed in leaves (16 mg GAE/g DW) when compared with roots (< 2 mg GAE/g DW), while NEPs presented a higher TPC in roots (21 mg GAE/g DW) than in leaves (13 mg GAE/g DW). EPs are the principal compounds present in leaves and NEPs in roots. The spectral fingerprint and fragmentation patterns of some C. chinense phenolic compounds were obtained by UPLC – PDA - MS. The analysis suggests that EPs detected in leaves (H1 – H3) are tentatively phenolics acids as chlorogenic and hydroxycinnamic acid derivates, and flavonoids (H4 – H7) as luteolin, quercetin, chrysoeriol or apigenin that provide protection against oxidative stress. The analysis suggest that NEPs compounds detected in roots (R1 – R3) are tentatively phenolic acids that are accumulated in cell wall and provide mechanical strength and protection against pathogen attack in roots.

Keywords. Phenolic compounds extraction, liquid chromatography, photodiode array detector, spectral fingerprint, mass spectrometry.

Resumen. La extracción de compuestos fenólicos solubles (CFS) y compuestos fenólicos no solubles (CFNS) en raíces y hojas de plántulas de Capsicum chinense fueron evaluadas usando extracción asistida por ultrasonido (UAE) y solventes con diferentes polaridades. Para los CFS en raíces y hojas de C. chinense no se encontraron diferencias significativas en el contenido de fenoles totales (CFT). Para los CFNS se observó un mayor CFT mediante extracción acida que con extracción alcalina. Para los CFS un mayor CFT se observó en hojas (16 mg EAG/g BS) que en raíces (< 2 mg EAG/g BS), mientras que para los CFNS un mayor CFT se observó en raíces (21 mg EAG/g BS) que en hojas (13 mg EAG/g BS) indicando que los CFS están principalmente distribuidos en hojas y los CFNS en raíces. Mediante UPLC – PDA- MS se obtuvo la huella espectral y el patrón de fragmentación de algunos compuestos fenólicos presentes en C. chinense. El análisis sugiere que los CFS detectados en hojas se tratan tentativamente ácidos fenólicos (H1 – H3) como derivados de ácido clorogénico y ácido hidroxicinámico, y flavonoides como luteolina, quercetina, chrysoeriol o apigenina, que tienen funciones como proveer protección frente el estrés oxidativo. Para los CFNS detectados en raíz (R1 – R3) el análisis sugiere que tentativamente se tratan de ácidos fenólicos acumulados en la pared celular con la función de proveer fuerza mecánica y protección frente al ataque de patógenos en las raíces.

Palabras clave. Extracción de compuestos fenólicos, cromatografía de líquidos, detector de matriz de fotodiodos, huella digital espectral, espectrometría de masas.



INTRODUCTION

The habanero pepper (Capsicum chinense) from Yucatan peninsula is a crop with great cultural and commercial importance1 that has a Protected Denomination Origin (PDO). This denomination distinguishes it from those produced in other regions.2 In order to obtain the fruit that comply with quality standards and productivity of the PDO (established in the NOM-189-SCFI-2017),3 it is necessary that the plant have an adequate physiological state and the capacity to adapt to constant environmental changes.4

The phenolic compounds are one of the main groups of secondary metabolites present in plants and these are characterized by the presence of a phenolic group in their structure. These compounds can be classified generally as flavonoids (flavones, flavonoids, anthocyanins, isoflavones, etc.) and non-flavonoids (phenolic acids, coumarins, stilbenes, lignins).5 In plants, phenolic compounds are related to the processes of environmental adaptation because they possess a wide range of biological and ecological functions. It can act as a pollinator attractor, herbivores and pathogens defense, and attract beneficial microorganisms from the rhizosphere. It is also known that these compounds facilitate seeds dispersion and protect them against UV irradiation and oxidation stress.6-9

Phenolic compounds are distributed in different tissues of the plant. A significant number of phenolic compounds are mainly stored in the cell vacuoles and can be easily extracted using an aqueous phase or organic solvents. Extractable phenolic compounds (EPs) are considered as the extractable fraction obtained using ethanol, acetone and methanol as solvent.5,10 Another important fraction of phenolic compounds is associated with macrostructures such as carbohydrates and proteins and is known as

non-extractable phenolics compounds (NEPs). The extraction of these fraction involves the use of acid or alkaline conditions that induce hydrolysis phenomena.10-12

A useful technique to separate and detect phenolic compounds is by performing ultra-high performance liquid chromatography (UPLC) paired with a photodiode array detector (PDA). Phenolic compounds are generally detected at a wavelength of 240 to 560 nm. Specifically, hydroxycinnamic acids are detected at 320 nm, flavones and flavonols between 350 – 365 nm, and anthocyanins between 460 – 560 nm. Furthermore, mass spectrometry (MS) is another effective technique to carry on structural characterizations and molecular mass determinations of phenolic compounds.9,10 After a purification process, more information about structural characteristics of these compounds can be obtained by using complementary techniques such as Fourier transformed infrared spectrometry (FTIR) or Nuclear Magnetic Resonance (NMR).13, 14

Extraction and identification of phenolics compounds is important because it contributes to the study of metabolic changes in plants against variation in the environment. In the case of Capsicum spp. an important stage occurs during seedling transplantation. Specifically, 45 days post-germination related to the increase in stress tolerance.15 Therefore, the main objective of this work was to study the effects of different EPs and NEPs extraction treatments on the total phenolic content (TPC) in habanero pepper roots and leaves. Moreover, the determination of extract profiles using UPLC – PDA/MS in order to know the distribution of phenolic compounds before transplantation as an important stage in the production of habanero pepper.



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Vegetal Material

C. chinense variety Chichen Itzá plants were obtained from a local greenhouse producer in Suma, Yucatán México (45 days post-germination).

Chemicals.

Chlorogenic acid, catechin, vanillic acid, ellagic acid, ferulic acid, m – coumaric acid and cinnamic acid. Solvents: HPLC grade acetone, methanol and water, 1- butanol (purity ≥ 94.4 %) and formic acid (purity ≥95 %) were purchased from Sigma-Aldrich.

Extraction methods

Capsicum chinense roots and leaves were frozen in liquid nitrogen and stored at - 40° C. The frozen fresh plant material was grounded (KRUPS, GX41000, MX) to obtain a powder with a particle size <500 µm. Samples (1 g) were placed in conical tubes (FalconTM) of 15 mL and homogenized in 10 mL of solvent.

Extraction of soluble phenolics compounds (EPs) in roots samples was carried out with ethanol 50 % (ER1), methanol (ER2), methanol 80 % (ER3) and acetone 80 % (ER4) as solvents. In the case of leaves samples, methanol (EL1), methanol 80 % (EL2) and acetone 80 % (EL3) were employed. Ultrasound-assisted extraction (UAE) was performed as extraction method at 80 % amplitude for 15 min according to Covarrubias-Cárdenas et al.16 In Table 1 shows specific conditions for phenolic extraction.

Extraction of non – extractable phenolic (NEPs) compounds a pre-extraction of EPs was performed according the above described using

pure methanol. Subsequently, the sample was centrifuged, and the supernatant was discarded. The sediment was washed with methanol to eliminate the residues of EPs contained in the sample (this process was repeated three times for each sample). NEPs extraction was performed in roots (ER6) and leaves (EL5) using acidic conditions (1mL HCl 2 N per 5 mL of butanol) and treated in a water bath at 90°C for 1 h (Table 1). Alkaline extraction was also performed in roots (ER5) and leaves (EL4) using NaOH (2N) and treated in a water bath at 60°C for 1 h. After acidic and alkaline extraction, pH was adjusted at pH 7 with NaOH (2N) or HCl (2N) respectively.10-12 Finally, all extracts were centrifuged at 6,500 rpm for 15 min and the supernatant was filtered with Millex Millipore filter (0.20 μm) and stored at - 40°C until further analysis.

Table 1. Techniques used in the extraction of phenolic compounds from roots and leaves of Capsicum chinense.

Sample(Code)

Phenolic Fraction Extraction conditions

Root(ER1) EPs

Ethanol: Water (50:50 v / v)Ultrasonication at 80% amplitude for 15 min,

Initial Temperature (25 °C)

Root(ER2) EPs

Methanol Ultrasonication at 80% amplitude for 15 min,


Root(ER3) EPs

Methanol: Water (80:20 v/v)Ultrasonication at 80% amplitude for 15 min,


Root(ER4) EPs

Acetone: Water (80:20 v/v)Ultrasonication at 80% amplitude for 15 min,

Initial Temperature (25 °C)Root(ER5) NEPs HCl (2N)/ Butanol

Extraction in water bath at 90 °C for 1 h Root(ER6) NEPs NaOH (2N)

Extraction in water bath at 60 °C for 1 h

Leaves(EL1) EPs

MethanolUltrasonication at 80% amplitude for 15 min,

Temp. Initial (25 °C)

Leaves(EL2) EPs

Methanol: Water (80:20 v/v)Ultrasonication at 80% amplitude for 15 min,


Leaves(EL3) EPs

Acetone: Water (80:20 v/v) Ultrasonication at 80% amplitude for 15 min,

Initial Temperature (25 °C)Leaves(EL4) NEPs HCl (2N)/ Butanol


Leaves(EL5) NEPs NaOH (2N)




Total phenolic content by Folin – Ciocalteu assay

The total phenolic compounds determination was performed by Folin – Ciocalteu assay using a spectrophotometer (Thermo Scientific, BIOMATE 3S, USA). TPC values were expressed as mg GAE g/ DW.16

Chromatography analysis by UPLC – PDA – MS

To identify the polyphenolic compounds, a mixture of polyphenolic standard from 10 to 100 ppm (chlorogenic acid, catechin, vanillic acid, ellagic acid, ferulic acid, m-coumaric acid, cinnamic acid) was prepared. Separation and detection of the standards and the analytes in the extracts were determined by UPLC analysis using Acquity H Class Mark Waters equipment paired with photodiode array detector (PDA) and mass spectrometry (MS).

Sample injection volume was 2 μL in a VanGuard Acquity UPLC HSS C18 precolumn and Acquity UPLC column at a flow rate of 0.2 μL / min. In the mobile phase, solution A of Water: HCOOH (99.9: 0.1) and solution B of MeCN: HCOOH (99.9: 0.1) was used. The PDA reading λ was performed in a range from 190 to 400 nm. The analytical response absorbance was taken at 290 nm. The samples were also analyzed with a compact quadrupole mass spectrometer (MS) Xevo TQ – S micro infused with electrospray ionization (ESI) with a flow of 10 μL / min. Analysis were performed principally in negative ion mode (some cases in positive ion mode) with an ionization voltage of 4,000 V and using N2 as a carrier gas.

Partial purification of extracted compounds

The phenolic compounds purification was performed using an Advanced Automated Flash Purification Biotage System (Model Isolera OneTM, Sweden). 1.5 mL of the sample was loaded

automatically on to a SNAP C18 30g Biotage cartridge. Elution was carried out at a flow rate of 25 ml/min using a gradient of (A) 0.1 % of formic acid in ultra-pure water and (B) 0.1 % formic acid in acetonitrile. The gradient was programmed as follows: 3 Column Volume (CV) of 0 % B isocratic; 1 CV linear gradient from 0 % to 10 % B; 1 CV linear gradient from 10 % to 23% B; 1 CV of 23 % B isocratic; 4 CV linear gradient from 23 % to 24 % B; 2 CV linear gradient from 24 % to 100 %, 3 CV from 100 % to 50 % B. Peak monitoring by UV detection was performed at a wavelength of 290 nm. Subsequently, the fractions obtained were analyzed by UPLC - PDA

Statistical analysis

TPC samples obtained in each treatment was expressed as the mean ± standard deviation of 3 determinations. A simple ANOVA was employed to perform statistical analysis. Tukey method was used for comparisons between the means of the treatments evaluated (p <0.05) using Statgraphics Centurion Version XVI software (Manugistic Inc., Rockville MD, USA).

RESULTS

Extractable phenolics (EPs) compounds in roots and leaves

EPs in roots extracts obtained by treatment ER1, ER2, ER3 and ER4 presented values of TPC of 1.97 ± 0.24, 1.57 ± 0.12, 1.87 ± 0.06 and 1.83 ± 1.06 mg GAE/ g DW, respectively. In leaves extracts obatined by treatment EL1, EL2 and EL3 the values of TPC obteined were 16.93 ± 2.16, 17.82 ± 2.64 and 17.30 ± 2.88 mg GAE/g DW, respectively. A higher TPC was found in EPs leaves extracts. Statistical analysis shows that there is no significant differences between treatments employed for the extraction of EPs in roots and leaves, respectively (Table 2).



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Non – extractable phenolic (NEPs) compounds in roots and leaves

NEPs in roots extracts obtained by treatment ER5 (acidic extraction) and ER6 (alkaline extraction) presented values of TPC of 21.46 ± 1.02 and 16.77 ± 0.56 mg GAE/g DW, respectively. In leaves extracts obtained by treatment EL4 (acidic treatment) and EL5 (alkaline extration) a TPC of 13.14 ± 0.37 and 11.25 ± 1.38 mg GAE/g DW was quantify. Higher values of TPC was found in NEPs roots. Statistical analysis shows that there is a significant difference between both treatments in roots and leaves. Furthemore, a higher TPC was found by acid extraction (Table 2).

Table 2. Total phenolic content (TPC) and number of phenolic compounds detected by UPLC – PDA in roots and leaves of Capsicum chinense using different extraction treatments.

Identified compounds in Capsicum chinense plants by UPLC - PDA

The chromatogram of the standards is shown in Figure 1. The order of elution of the polyphenol standards was as follows: chlorogenic acid (A) < catechin (B) < vanillic acid (C) < Ellagic acid (D) ferulic acid (E) < coumaric acid (F) < cinnamic acid (G).

The time retention (RT) and maximum peaks (λmax) in UV light fingerprint (190 to 400 nm) of de phenolics compounds standards and analytes in roots (R1 – R3) and leaves (H1 – H7) of Capsicum chinense are shown in Table 3.

Root extracts obtained by treatments coded as ER1, ER2, ER3 and ER4 did not presented compounds (Table 2). Nevertheless, it was possible to observe the presence of some peaks in ER5 (Figure 2) and ER6 (Figure 3) after using alkaline and acid hydrolysis conditions. In ER5 sample, a compound coded as R1 (RT: 6.71 min, 276 nm) was detected, while in ER6 a compound code as R3 (RT: 8.65 min, λmax: 217, 326 nm) was detected. Data shown in Table 3.

In leaves extracts the presence of seven peaks were detected from treatments coded as EL1, EL2 and EL3 (Figure 4). The compounds were coded as H1 (RT: 8.25 min, λmax: 193, 217, 279 nm), H2 (RT: 9.13 min, λmax: 210, 311 nm), H3 (RT: 9.71 min, λmax: 193, 211, 308 nm), H4 (RT: 10.17 min, λmax: 206, 254, 349 nm), H5 (RT: 11.35 min, λmax: 198, 266, 337 nm), H6 (RT: 11.66 min, λmax: 196, 252, 266, 348 nm) and H7 (RT: 14.82 min, λmax: 211, 266, 337 nm) and these are shown in Table 3. Using acid and alkaline hydrolysis conditions in treatment EL4 and EL5 none compound was identified (Table 2).

Fig. 1. Separation of phenolic compounds standards by UPLC – PDA (Analytical response at 290 nm). The elution order of the compounds is shown as A: chlorogenic acid, B: catechin, C: vanillic acid, D: Ellagic acid, E: ferulic acid, F: coumaric acid, G: cinnamic acid.

Treatment Sample Phenolic Fraction

TPC(mg GAE/g DW)

Compounds detected by UPLC-PDA

(ER1) Roots EPs 1.97 ± 0.24 0

(ER2) Roots EPs 1.57 ± 0.12 0

(ER3) Roots EPs 1.87 ± 0.06 0

(ER4) Roots EPs 1.83 ± 1.06 0

(ER5) Roots NEPs 21.46 ±1.02 2

(ER6) Roots NEPs 16.77 ± 0.56 1

(EL1) Leaves EPs 16.93 ± 2.16 7

(EL2) Leaves EPs 17.82 ± 2.64 7

(EL3) Leaves EPs 17.30 ± 2.88 7

(EL4) Leaves NEPs 13.14 ± 0.37 0

(EL5) Leaves NEPs 11.25 ± 1.38 0

AU

Time (min)

A

BD

EF

G

C



Fig. 2. Detection of phenolic compounds in extract root (ER5 obtained by acidic extraction) of Capsicum chinense by UPLC – PDA. Two compounds (R1 and R2) were detected with analytical response at 290 nm.

Fig. 3. Detection of phenolic compounds in extract root (ER6 obtained by alkaline extraction) of Capsicum chinense by UPLC – PDA. A compound (R3) were detected with analytical response at 290 nm.

Fig. 4. Separation and detection of phenolic compounds in extract leave (EL2) of Capsicum chinense by UPLC – PDA. Analytical response at 290 nm.

Analysis of compounds in roots and leaves extracts by mass spectrometry

Fragmentation pattern for analytical standards are shown in Table 3. In root extract fragmentation, patterns for the two compounds were obtained. The fragment at m/z 109, 89, 53, 79, and 127 was found in R1 and fragments at m/z 217, 79, 117, 59 and 119 in R2 that are presented in descending order of relative abundance (Table 4). In leaves extracts, the fragmentation pattern for the three compounds was also obtained. Fragments at m/z 203, 327, 285, 116 and 204 in H1, fragments at m/z 337, 191, 177, 338, and 329 m / z in H2, and fragments at m/z

191, 337, 359, 341 and 371 m / z in H3 were found and presented in descending order of relative abundance (Table 3).

Table 3. Phenolic compounds: analytical standards and analytes detected in roots and leaves extracts of Capsicum chinense. Retention time (RT) and maximum peaks (λmax) in fingerprint UV light (190 to 400 nm) of the phenolics compounds standards and analytes in roots and leaves are shown. Also, fragmentation pattern and their relative abundance for two and three compounds in roots (R1 and R2) and leaves (H1 – H3) respectively were obtained.

RT(min) Compound λ Max

(190 to 400 nm)

Fragmentation patternm/z (% relative abun-

dance)

8.43 Chlorogenic acid (A) 216, 326 191 (100), 353 (95)

8.53 Catechin(B) 203, 279 289 (100), 179 (5)

8.92 Vanillic acid(C) 218, 260, 292 135 (100), 152 (7),

167 (5)

9.71 Ellagic acid(D) 219, 253 301 (100)

10.30 Ferulic acid (E) 194, 218, 322 133 (100), 193 (65), 178 (55), 149 (15)

10.86 m – coumaric acid (F) 195, 215, 277 119 (100), 163 (58),

18.76 Cinnamic acid (G) 217, 276 147 (100), 117 (62),

103 (45), 135 (25)

6.71 R1 193. 284*109 (100), 89 (58),

53 (24),79 (12), 127 (12)

6.89 R2 209, 276*217 (100), 79 (47),

117 (27), 59 (24), 119 (20)

8.65 R3 217, 326 -

8.25 H1 193, 217, 279203 (100), 327 (42), 285 (39), 116 (20),

204 (19)

9.13 H2 210, 311337 (100), 191 (88), 177 (19), 338 (17),

329 (8)

9.71 H3 193, 211, 308191 (100), 337 (85), 359 (10), 341 (5),

371 (3)10.14 H4 206, 254, 268, 349 -11.35 H5 198, 266, 337 -11.66 H6 196, 252, 268, 348 -14.32 H7 211, 266, 337 -

Information of fragmentation pattern was used to search for coincidences of compounds detected in C. chinense roots an leaves with those reported in spectral database of organic compounds (SDBS) and MassBank (https://massbank.eu/MassBank/Search). However, no match was found.

Time (min)

H5

AU H1H2

H3

H4H6

H7

1.00 2.00 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 11.00 12.00 13.00 14.00 15.00 16.00 17.00 18.00 19.000.0

2.5e-2

5.0e-2

7.5e-2

1.0e-1

1.25e-1

1.5e-1

1.75e-1

2.0e-1

2.25e-1

2.5e-1

2.75e-1

R1

R2AU

Time (min)

Time (min)

AU

R3



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After peak identification of phenolics compounds in roots and leaves extracted by UPLC – PDA, a partial purification was performed. Using the advanced purification system, partial separation of some compounds found in root and leaves was achieved from the extracts ER6 and EL2 that showed a higher response intensity in the chromatographic profiles. A total of 28 fractions was obtained for both samples.

In ER6 root extract partial purification of the compound R3 was possible and fraction 8 was obtained (Figure 5). In EL2 leaves extract, fraction 11 was obtained from the partial purification of H4 and H5 compounds (Figure 6). This presence was confirmed based on the retention time and the maximum peaks found in the spectral UV light fingerprint obtained by UPLC – PDA.

Fig. 5. Separation of phenolic compounds in roots extract of Capsicum chinense using an Advanced Automated Flash Purification Biotage System. Compounds R3 was partially purified in fraction 8.

Fig. 6. Separation of phenolic compounds in leaves extract of Capsicum chinense using an Advanced Automated Flash Purification Biotage System. Compounds H4 and H5 were partially purified in fraction 11.

DISCUSSION

Extractable phenolics compounds (EPs) in root and leaves

A low response of TPC (< 2 mg GAE/g DW) in treatment ER1, ER2, ER3 and ER4 can be explained probably because secondary metabolites such as phenolic compounds in roots are only produced under stress conditions. These compounds are exuded to modify the rhizosphere and have function as attractants of beneficial microorganisms from the soil, and defense mechanism against pathogen attack.8 It is possible that the development stage of the plant or stress absence in seedlings has an influence on the content of phenolic compounds in the roots.In treatments EL1, EL2 and EL3 a high response of TPC was found for EPs in leaves (16 mg GAE/g DW approximately). Therefore, the three solvents used are an adequate option to extract phenolic compounds from C. chinense leaves. A higher TPC concentration in leaves than roots probably because to phenolics compounds in C. chinense have an important function against UV light, drought and oxidative stress. Rodríguez-Calzada et al. reported that in C. annuum the UV light influences the accumulation of phenolic compounds such as chlorogenic acid and apigenin,17 while, Okunlola et al. reported that in C. chinense drought stress produces phenolic compounds accumulation.18

Non-extractable phenolics (NEPs) in roots and leaves

Results suggest that acid and alkaline extraction release a great quantity of phenolic compounds from macrostructures present in roots (21.46 ± 1.02 and 16.77 ± 0.56 mg GAE/g DW, respectively) and in leaves (13.14 ± 0.37 and 11.25 ± 1.38 mg GAE/g DW, respectively). These phenolics compounds are covalently

Time (min)

AU Fraction 8

Time (min)

AU Fraction 11



bound to structural components in cell wall and have functions as chemical and physical barriers against pathogens, animals and insects attack.9

A higher content of NEPs was found by performing acid extraction in roots and leaves. However, several authors have reported that a higher TPC is recovery by alkaline extraction than acid. This might be due to the appropriately rupture of the ester bond linking the phenolic acid to the cell wall polysaccharides. In addition, acid extraction at high temperatures causes degradation of the phenolic compounds. The acid extraction breaks glycosidic bonds and solubilizes sugars, but generally it has no effect on ester bonds.12 It suggests that a higher TPC in leaves and roots. might be because a great quantity of phenolic compounds bound to the cell wall by glycosidic bounds.

Compound Idetification in Capsicum chinense plants by UPLC – PDA and mass espectrometry analysis.

Roots extracts obtained by treatment ER1, ER2, ER3 and ER4 did not presented chromatographic peaks due to the low concentration of TPC (< 2 mg GAE/g DW) (Table 2). EPs in roots probably are exudate only after transplantation in C. chinense seedlings due to the adaptive function against possible stress factors in soil.8 In roots extracts obtained by treatment coded as ER5 (acid extraction) the compounds R1 and R2 were detected (Table 2). These compounds presented a spectral fingerprint characteristic to phenolics acids. Fragmentation pattern suggest that both compounds have a low molecular weight and present a possible molecular ion at m/z 109 and m/z 217 respectively. In treatment coded as ER6 (alkaline extraction), the compound R3 was detected and tentatively identified as chlorogenic acid because it presents similarity

in time retention and spectral fingerprint of the analytical standard (Table 3). According to the results some authors reported that different phenolic acids are bound to cell wall to increase mechanical strength that could be a possible function of phenolics compounds in C. chinense roots.9, 12

Seven chromatographic peaks (H1 – H7) in leaves extracts were detected in treatments EL1, EL2 and EL3 indicated that all methods are suitable for extraction of phenolics compounds from C. chinense leaves. Results suggest that compounds H1 to H3 are phenolic acids and H4 to H7 are flavonoids according to spectral fingerprint (Table 3). Only peak H3 presented the same retention time that ellagic acid. However, it did not show any match with the spectral finger print. Peak H1 presented similar fingerprint to some phenolic acids like cinnamic and m-coumaric acid, and possible molecular ion at m/z 203 or 337 due to its relative abundance. Compounds H2 and H3 presented similar fingerprint to chlorogenic acid and fragment at m/z 191 that suggests presence of quinic acid in the structure (Table 3).19

These results suggest that probably some phenolic acids in C. chinense leaves are chlorogenic or hydroxycinnamic acid derivates. Presence of chlorogenic and hydroxycinnamic acids are reported in C. annuum fruits. Tentatively, peaks H4 and H6 are luteolin, quercetin or chrysoeriol derivates, and peaks H5 and H7 tentatively are apigenin derivates according to the fingerprints reported by Morales-Soto et al. found in C. annuum fruits.19 These flavonoids in plants are primarily reported as compounds that reduce reactive oxygen species (ROS) produced by UV – B irradiance6,7,18 and particularly, apigenin have this adaptative function against UV stress. In leaves extracts obtained by treatment EL4 (acid extraction) and EL5 (alkaline extraction) no chromatographic peaks were observed (Table 2).



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In order to confirm the identification of the compounds, the use of purification procedures is required. It is important to use complementary techniques like nuclear magnetic resonance (NMR) or Fourier transform infrared spectrometry (FTIR) to reveal more information that allows structural and functional analysis of phenolics acids and flavonoids present in C. chinense plants.13, 14

CONCLUSIONS

EPs extraction was carried out by using different solvents but it did not show a significant effect in TPC of roots and leaves extracts. A higher TPC was found in leaves than in roots extracts possibly due to phenolic compounds accumulation if leaves generated to protect the plant against UV and oxidative stress. In the vase of NEPs, significant differences were observed in TPC of acidic and alkaline extractions. A higher TPC was obtained by acid extraction possibly due to phenolics compounds in roots and leaves present glycosidic bonds with macrostructures in the cell wall. A higher concentration of TPC was observed in roots than in leaves extract, probably because phenolics compounds provide mechanical strength in roots. A higher EPs content is mainly distributed in leaves, while NEPs are presented in roots of C. chinense plants. UPLC – PDA – MS analysis indicated that EPs presented in leaves are tentatively phenolics acid ( H1 – H3) as chlorogenic and hydroxycinnamic acid derivates, and some possible flavonoids (H4 – H7) like luteolin, quercetin, chrysoeriol and apigenin derivates. These have the function of reducing oxidative stress, while NEPs in roots probably are low molecular weight phenolics acids (R1 – R3) that provide mechanical strength in the cell wall. Partial purification of phenolics compounds H4 and H5 from EL2 extract and

R3 presented in ER6 extract was confirmed by UPLC-PDA. This allowed a more specific characterization of these molecules through complementary techniques.

FUNDING AND ACKNOWLEDGMENTS

To the project frontiers of science (No. 35) called: “Metabolic changes and the signal transduction system associated with the interaction of Capsicum chinense with Pythium sp.”, for the resources granted.

To CONACYT to Emanuel Herrera grant awarded.

To ASPELAB ® to provide de equipment (Advanced Automated Flash Purification Biotage System, Model Isolera OneTM, Sweden) for partial purification performed in this work.

Jamie Prutt, peace corps volunteer to the final English editing.

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7) León-Chan, RG, López-Meyer M, Osuna-Enciso T, Sañudo-Barajas JA, Basilio-Heredia J, León-Félix J. 2017. Low temperature and ultraviolet – B radiation affect chlorophyll content and induce the accumulation of UV-B-absorbing and antioxidant Compounds in bell pepper (Capsicum annuum) plants. Environ. Exper. Bot. 139: 143 – 151. doi: 10.1016/j.envexpbot.2017.05.006

8) Badri DV, Vivanco JM. 2009. Regulation and function of root exudates. Plant Cell Environ. 32: 666 – 681. doi: 10.1111/j.1365-3040.2008.01926.x.

9) Jean-Christophe C, Casas MI, Yang F, Grotewold E, Alonso AP. 2019. Beyond the wall: High-throughput quantification of plant soluble and cell-wall bound phenolics by liquid chromatography tandem mass spectrometry. J. Chromatogr. A. 1589: 93 – 104. doi: 10.1016/j.chroma.2018.12.059

10) Domínguez-Rodríguez G, Marina ML, Plaza M. 2017. Strategies for the extraction and analysis of non-extractable polyphenols from plants. J Chromatogr A. 1514: 1 – 15.

doi: 10.1016/j.chroma.2017.07.06611) Guan-Lin C, Zhang X, Song-Gen C, Men-

Di H, Yong-Qing G. 2017. Antioxidant activities and contents of free, esterified and insoluble-bound phenolics in 14 subtropical fruit leaves collected from south of China. J. Funct. Foods. 30: 290-302. doi: 10.1016/j.jff.2017.01.011

12) Acosta-Estrada BA, Gutiérrez-Uribe JA. Serna-Saldívar SO. 2013. Bound phenolics in foods, a review. Food Chem. 152: 46-55. doi: 10.1016/j.foodchem.2013.11.093

13) Materska M, Piacente S, Stochmal A, Pizza C, Oleszek W, Perucka I. 2003. Isolation and structure elucidation of flavonoid and phenolic acid glycosides from pericarp of hot pepper fruit Capsicum annum L. Phytochem. 63: 893 – 898. doi: 10.1016/S0031-9422(03)00282-6

14) Woo-Ri K, Eun OK, Kyungsu K, Oidovsambuu S, Sang HJ, Byung S, Chu WN, Byung-Hum U. 2014. Antioxidant Activity of Phenolics in Leaves of Three Red Pepper (Capsicum annuum) Cultivars. Agricultural and Food Chem. 62 (4): 850-859. doi: 10.1021/jf403006c

15) Vázquez-Casarrubias G, Escalante-Estrada JAS, Rodríguez-González MT, Ramírez-Ayala C, Escalante-Estrada LE. 2011. Age at transplant and its effect on growth and yield of chilli apaxtleco. Rev. Chapingo Ser. Hortic. 12 (1): 61-65. doi: 10.5154/r.rchsh.2011.17.009

16) Covarrubias-Cárdenas AG, Martínez-Castillo JI, Medina-Torres N, Ayora-Talavera T, Espinosa-Andrews H, García-Cruz NU, Pacheco N. 2018. Antioxidant Capacity and UPLC – PDA – ESI – MS Phenolic Profile of Stevia rebaudiana Dry Powder Extracts Obtained by Ultrasound Assisted Extraction. Agron. J. 8 (9): 170. doi: 10.3390/agronomy8090170

17) Rodríguez-Calzada T, Qian M, Strid A, Neugart S, Schreiner M, Torre-Pacheco



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I, Guevara-González RG. 2019. Effect of UV-B radiation on morphology phenolic compound production, gene expression, and subsequent drought stress responses in chili pepper (Capsicum annuum L.). Plant Physiol. Biochem. 134: 94-102. doi: 10.1016/j.plaphy.2018.06.025

18) Okunlola GO, Olatunji OA, Akinwale RO, Tariq A, Adelusi AA. 2017. Physiological response of three most cultivated pepper species (Capsicum spp.) in Africa to drought stress imposed at three stages of growth and development. Sci. Hort. 224: 198-205. Doi: 10.1016/j.scientia.2017.06.020

19) Morales – Soto A, Gómez – Caravaca A M, García – Salas P, Segura – Carretero A, Fermández – Gutiérrez A. 2013. High-performance liquid chromatography coupled to diode array and electrospray time-of-flight mass spectrometry detectors for a comprehensive characterization of phenolic and other jj j j polar compounds in three pepper (Capsicum annuum L.) samples. Food Res. Int. 51: 977 – 984. doi: 10.1016/j.foodres.2013.02.022



Research article

Chemical stability and residual plasmin activity of milk subjected to various pressure-temperature treatmentsParada-Rabell Francisco1, Chavez-Parga Maria del Carmen2, Alvarez B. Valente*3 1 CytoSport Inc. a division of Hormel Foods Corporation, Walnut Creek, California, USA. 2 Facultad de Ingenieria Quimica, Universidad Michoacana de San Nicolas Hidalgo, Morelia, Michoacan, Mexico.3Department of Food Science, The Ohio State University, Columbus, Ohio, USA.


Received: 12 november 2018/ Accepted: 26 march 2019/ Published online: 31 may 2019

Abstract. The effects of different pressure-temperature combinations on the chemical stability and residual plasmin activity in milk were studied and compared to those of pasteurized (77 ± 0.8oC for 18 s) and ultra-high temperature (UHT; 138 ± 1oC for 2 s) processed milk. Milk was pressure-assisted thermal processed (PATP) using a factorial 3x1x3 model at temperature (32, 78 and 105oC), pressure (650 MPa), and time (0, 1, and 5 min). The chemical stability of milk samples was evaluated based on its proteolysis values over a period of 90 d under different storage conditions. Total plasmin activity was analyzed after extraction from milk samples using BODIPY FL-Casein as a substrate. No significant proteolysis was observed in pressure-treated samples at the end of their respective shelf life. Plasmin inactivation rates increased with increasing temperature during pressurization. Although the enzyme was not completely inactivated, combinations of temperature (105oC) with ultra-high pressure (650 MPa) were sufficient to decrease the enzyme activity to levels comparable to UHT processes. These results suggest that the pressure–temperature combinations used in this study were capable of rendering milk with enzymatic activity close to that of pasteurized and UHT milk. However, processing temperature and storage conditions had a significant role in the extent of protease activity in milk.

Keywords. Pressure-Assisted Thermal Processing, proteolysis, lipase, dairy products

Resumen. Los efectos de diferentes combinaciones de presión y temperatura en la estabilidad química y actividad residual de plasmina en leche fueron evaluados y comparados con aquellos de leche pasteurizada (77 ± 0.8oC por 18 s) y leche UHT (ultra-high temperature, por sus siglas en inglés; 138 ± 1oC por 2 s). La leche fue procesada con alta presión-temperatura utilizando un modelo factorial 3x1x3 a temperatura (32, 78 y 105oC), presión (650 MPa), y tiempo (0, 1, y 5 min). La estabilidad química de las muestras de leche fue evaluada en base a sus valores de proteólisis durante 90 d bajo diferentes condiciones de almacenamiento. La actividad total de Plasmina fue analizada después de extraerla de la leche utilizando BODIPY FL-Casein como sustrato. No se observó proteólisis significativa en la leche procesada con diferentes presiones y temperatura al final de su vida de anaquel. La proporción de Plasmina inactiva en la leche fue mayor conforme aumentó la temperatura durante la presurización. Aunque la enzima no fue completamente inactivada, combinaciones de presión (650 MPa) con temperatura (105

oC) fueron suficientes para disminuir la actividad enzimática a niveles similares a los del proceso UHT. Nuestros resultados sugieren que las combinaciones de presión y temperatura utilizados en este estudio fueron capaces de reducir la actividad enzimática de la leche a niveles cercanos a los de la leche pasteurizada y de la leche UHT. Sin embargo, la temperatura de proceso y las condiciones de almacenamiento afectan la actividad de proteasas en la leche.

Palabras clave. Procesamiento térmico asistido por presión, proteolisis, lipasa, productos lácteos



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INTRODUCTION

Chemical changes due to enzymatic activity are not desirable in fluid milk as they contribute to adverse organoleptic effects and limit its shelf life. In addition to bacterial enzymes, the activity of up to seventy naturally occurring or indigenous enzymes has been identified in fluid milk and only twenty have been fully characterized.1 Lipoprotein lipase (LPL) and plasmin are probably the most significant enzymes responsible for deterioration of milk constituents due to their high temperature stability.1,2,3 Plasmin (EC 3.4.21.7) is the principal indigenous proteinase in milk.4,5,1,6 The enzyme is responsible for proteolysis of the Lys and Arg peptide bonds of as1-, as2- and β-caseins, and it has optimal activity values at pH 7.6 and 37oC.7,1,5 The enzyme is an alkaline trypsin-like proteinase that consists of a complex system formed by plasmin, its inactive form plasminogen, plasmin/plasminogen inhibitors and plasminogen activators (PAs). Upon storage of milk, plasminogen is converted into active plasmin by cleavage of the Arg557–Ile558 bond by two types of proteinases: urokinase and tissue-type Pas.4,1 Therefore, higher plasmin activity values are observed in pasteurized milk upon storage.5 Plasmin is also a heat resistant enzyme. Temperatures up to 120oC for 15 min, are necessary to ensure absence of plasmin activity in milk.3

Emerging technologies, such as high pressure processing (HPP), have been developed in recent decades to process foods, inactivate bacteria and their enzymes with less thermal damage. Data available on inactivation of proteases by HPP is fairly limited.8,9,10,11,12 Researchers suggest that the application of high pressure in combination with temperature seems an excellent alternative to traditional thermal treatments to inactivate heat resistant enzymes in milk. By choosing adequate process combinations of high pressure and temperature, bacterial spores and their enzymes

can be inactivated without the application of excessive heat and without compromising the quality characteristics of milk.13,14,15 Therefore, the application of high temperature (90-121oC) in combination with high pressure (500-700 MPa) could significantly extend the shelf life of milk. The objectives in this study were to evaluate the chemical and enzymatic stability of high-pressure processed (HPP) and pressure-assisted thermal processed (PATP) milk. Proteolysis in these samples was compared against that found in commercial high-temperature-short-time (HTST) pasteurized and ultra-high temperature (UHT) processed milk over time used as controls. Additionally, a sensitive protease assay was developed to measure residual plasmin activity concentration in milk samples.


Milk preparation

Raw milk was obtained from a commercial dairy plant in Ohio (Orrville, OH) and transported to the OSU dairy pilot plant. The temperature of milk was kept at ~4oC at all times during transportation (approximately 2 h). Milk was standardized to 2% milkfat and two-stage homogenized (500/1000 psi) using a Lab 100 M-G homogenizer (Lubeck-Schlutut, Germany). Homogenized milk was immediately cooled to 7 ± 1oC and stored under refrigeration conditions (4 ± 1oC) until HPP and PATP processing. All milk samples were subjected to various pressure-heat processing conditions within 72 hrs. On the day of processing, milk samples were preheated to their corresponding initial temperature (Ti) using an UHT/HTST Lab-25HV Hybrid unit (Micro Thermics Inc., Raleigh, NC). The initial temperature of milk samples was adjusted as a function of the final target pressure during processing. This temperature was estimated based on the heat of compression of water and skim milk for various temperatures.16 The preheating time for milk to achieve the desired initial



temperature was 2.5 min. The temperature of milk samples during preheating was set at values slightly higher than their required initial temperature (Ti) to compensate for the heat loss that occurred between filling and high pressure processing. The initial temperature of milk samples was estimated based on the following equation:16

Equation 1: where, Ti = initial temperature before pressurization, Tmax = final processing temperature during pressurization, CH = heat of compression, and DP = applied pressure, DTH = Heat loss during pressurization.

Ti accounted for heat loss during preheating until pressure treatment. The preheating temperature of milk samples was set at 6, 53 and 78 ± 1oC, respectively, depending on the final pressure-heat treatment applied. Milk was filled into light-protected 8 oz polyethylene teraphtalane (PET) bottles without head space and manually capped. Immediately after capping, milk samples were placed in a water bath set at the same temperature as the milk and then high pressure processed. There was a minimal heat loss of 1 ± 0.5oC during this step.

High pressure processing

An S-IL-110-625-08-W cold isostatic press system (Stansted Fluid Power Ltd., Essex, UK) was used to high pressure process (HPP; 650 MPa, 32 and 78oC) and pressure-assisted thermal process (PATP; 650 MPa, 105oC) milk samples. A 1:1 ratio propylene glycol to water mixture was used as the pressure-transmitting fluid. Milk samples were processed using a factorial 3x1x3 model at temperature (32, 78, and 105oC), pressure (650 MPa) and time (0, 1 and 5 min) in duplicate. Pressure come-up time ranged between 1.5 to 3 min depending on the treatment applied, and decompression time was 1.2 min. The temperature of the vessel and pressure-transmitting

fluid was thermostatically adjusted to the desired initial processing temperature by circulating propylene glycol through the external jacket of the pressure chamber. This temperature was set at the same initial temperature as the milk samples (4, 48 and 73oC, respectively). Milk samples were filled into a cylindrical sample basket (102 mm dia x 559 mm height) (Stansted Fluid Power Ltd., Essex, UK) and loaded into high-pressure equipment using a mechanical lift mechanism. The temperature of the milk samples during various pressure treatments was monitored at the center of the carrier basket using a T-type thermocouple (Omega engineering, CT, USA), which was placed inside a milk sample bottle designated for temperature recording purposes. After decompression, pressure treated milk bottles were immediately placed in an ice-water bath to stop further thermal effect, and stored at either room temperature (25 ± 1oC) or refrigeration (4 ± 1oC) conditions until analyzed for their proteolysis values over a period of 20, 60 and 90 d, depending on the final pressure-temperature treatment applied. Extra aliquots (n = 4) of each milk sample were taken on 0 d and quick frozen (-40oC) until analyzed for their total plasmin activity.

Thermal treatments

HTST pasteurized milk (2% milkfat), obtained from the same batch of raw milk used for pressure-heat treatments, was processed in a commercial dairy operation (Orrville, OH) at 77 ± 0.8oC for 18 s. Additionally, UHT treated (138 ± 1oC for 2 s) milk samples were obtained commercially (Wallington, NJ). HTST pasteurized and UHT milk samples were stored under refrigeration conditions (4 ± 1oC) and room temperature conditions (25 ± 1oC), respectively.

Milk analyses

The extent of proteolysis and residual plasmin activity in HPP, PATP, HTST and UHT milk samples was investigated. Due to equipment



31

limitations, milk samples were obtained from two different sources and variations in the chemical stability and residual plasmin activity of milk samples were expected. However, the intent of this study was to evaluate the effect of different combinations of pressure-heat treatments on the overall enzymatic activity of milk samples. Milk was exposed to similar temperature histories, with or without pressure, comparable to traditional thermal treatments. And, HTST pasteurized (77 ± 0.8oC for 18 s) and UHT (138 ± 1oC for 2 s) milk samples were chosen as controls. Milk samples were taken at regular intervals over a period of 20, 60 and 90 d as shown in Table 1. The effect of processing on the residual plasmin activity of milk was measured on the initial day of analysis (0 d).

Table 1. Longitudinal scheme of milk analyses

aGroup

Time (days)b

0 2 4 6 10 15 20 30 45 60 75 90

A ● ● ● ● ● ● ●

B ● ● ● ● ● ● ● ●

C ● ● ● ● ● ● ●

a Group A: HTST pasteurized (77±1oC for 18 s) and High Pressure Processed (HPP) milk (650 MPa, 32oC for 0, 1 and 5 min, respectively). Group B: HPP milk (650 MPa, 78oC for 0, 1 and 5 min). Group C: Ultra High Temperature processed (UHT) (138±1oC for 2 s) and Pressure Assisted Thermal Processed (PATP) milk (650 MPa 105oC for 0, 1 and 5 min)b Denotes day of analysis.

Proteolysis

Proteolysis in milk samples was evaluated using sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) analysis according to the method described by Verdi et al.,17 with modifications. Milk (2 mL) was centrifuged (4600 rpm, 15 min) to separate milkfat before electrophoresis was run. Skim milk was diluted 1:10 (v/v) in sample buffer (35.5% deionized

water, 62.5 mM Tris-HCl pH 6.8, 20% glycerol, 2% sodium dodecyl sulfate (SDS), 0.01% bromophenol blue and 0.5% β-mercaptoethanol), and vials were immersed in boiling water for 4 min. Aliquots (8.5 µL) of milk samples were then loaded into 12% resolving Tris-HCl gels (Bio-Rad Laboratories Inc., Hercules, CA). Electrophoresis was conducted at a constant current of 145 V using a 2-gel, 10-well, 1.0 mm thickness vertical electrophoresis system (Bio-Rad Mini-Protean Tetra Cell Hercules, CA) until dye reached the bottom edge of the gel (approximately 2 h).

Gels were maintained at 4 ± 1oC during the electrophoresis run. Subsequently, gels were washed three times with deionized water and copper-stained with 0.3 M CuCl2 (Fisher Science Education, Rochester, NY).18 A broad range pre-stained SDS-PAGE standard (Bio-Rad Laboratories Inc., Hercules, CA) was run along milk samples to identify electrophoretic bands based on their molecular weight. Gels were scanned using a densitometer (Bio-Rad GS-800, Hercules, CA). The proportion of proteolysis products formed as a result of protease activity was determined by measuring the absolute decrease in area of known casein bands (as-casein, β-casein, and κ-casein). This decrease in area was proportional to the total increase in area of bands identified as proteolysis products, as shown in Figure 1. Lanes 1-6 show the change in known protein bands that occurred over the course of 15 days in milk samples processed at 650 MPa, 32oC for 5 min and stored under refrigeration. Any increase of intensity in electrophoretic bands (proteolysis products) was also considered an index of proteolysis. Results were expressed as the sum of relative percentage of proteolysis of casein bands (intact minus enzymatically damaged) at the end of the shelf life of milk samples and according to the following equation:17



Equation 2: where, (Int x mm2)f is the intensity x area of casein electrophoretic bands at the end of the shelf life of milk. And, (Int x mm2)0 is the intensity x area of casein electrophoretic bands on 0 d.

Fig. 1 SDS-PAGE electrophotogram of milk subjected to High Pressure Process treatment. (650 MPa, 32oC for 5 min). *Lane 1-6 milk sampled on 0, 2, 4, 6, 5, 10 and 15 days after stored at 4±1oC.**Unk 1 – 3: major casein proteolysis products identified as unknown

Plasmin Activity

Residual plasmin activity was evaluated in pressure-treated milk samples and the controls HTST pasteurized and UHT samples. A sensitive spectrofluorometric assay was developed using 4, 4-Difluoro-5, 7–dimethyl -4-bora-3a, 4a–diaza-s-indacene-3-propionic acid, succinimidyl ester (BODIPY FL, SE)-labeled casein (Invitrogen Corp., Carlsbad, CA) as a fluorogenic substrate for plasmin.19 This substrate is cleaved by a wide variety of proteases, including plasmin, into small fluorescent peptides in a linear time-dependent manner during the enzymatic reaction. The resulting increase in fluorescence upon hydrolysis is proportional to enzymatic activity. Unlike other spectrofluorometric assays, the use of BODIPY FL-Casein did not involve any separation steps; and allowed for a direct, continuous fluorescent measurement upon incubation with plasmin. BODIPY FL-

Casein belongs to a family of boron-containing flourophores with emission spectra between 510 to 610 nm.19,20

The substrate has extinction coefficients near 80 000 M-1 cm-1, and it resists changes in the ionic strength of the sample over a wide range of pH. A disadvantage of the substrate is its very small Stoke’s shift, and therefore a higher tendency to shelf- quenching with increasing protein labeling.20 BODIPY FL labeled casein has approximately 5 mole of dye/mole of protein,21 and the highest fluorescence intensity values were observed using a substrate solution of 10 mg/ml (data not shown).

A standard curve was constructed with known concentrations (0, 0.5, 1, 1.5, 2 and 3 mg/mL) of purified plasmin (Sigma-Aldrich, St. Louis, MO) in phosphate buffer, pH 7.6. A 10 mg/mL substrate solution (BODIPY FL-Casein) was also prepared in 20 mM phosphate buffer, pH 7.6 + 150 mM NaCl. The range of enzyme response was determined by mixing equal aliquots (375 µL) of each enzyme dilution with BODIPY FL-Casein substrate solution. Samples were incubated for 1 h at 37oC, protected from light. Fluorescence intensity was measured in 1-cm path length cuvettes at 485 nm excitation and 530 nm emission filters, slits opened at 10/10, 1 s average reading time and voltage between 590-610 V using a Varian Cary Eclipse Fluorescence spectrophotometer (Walnut Creek, CA). Fluorescence emission intensities versus enzyme concentration were plotted; and limits of detection were determined (n = 4) using regression analysis (y = 52615x + 106.23; R2 = 0.997). Since equal amounts (375 µL) of substrate and enzyme solution were added to the mixture, a 1:1 (v/v) dilution ratio was considered in the final calculations.

Total plasmin activity was determined in milk samples after extraction of plasmin and plasminogen fractions.22,7 Milk (5 mL) was centrifuged at 4600 rpm for 15 min and the

{ } [ ] ( )( )

×

×−=

02

2

mmIntmmInt

100100(%) sProteolysi Relative fx



33

cream layer on the surface was discarded. Skim milk was then centrifuged at 32700 rpm for 1 h at 4oC. The supernatant (whey fraction) was discarded,; and the pellet (casein fraction) was reconstituted in 20 mM phosphate buffer, pH 7.6 + 50 mM ε-aminocaproic acid (EACA) (Sigma-Aldrich, St. Louis, MO). The mixture was incubated for 2 h at room temperature (25 ± 1oC), and then centrifuged at 32700 rpm for 1 h at 4oC to allow dissociation and transfer of the plasmin system from the casein micelle into the buffer. Plasmin and plasminogen content was determined after activation with 1600 IU/mg of urokinase (American Diagnostica Inc., Stamford, CT). Equal aliquots (0.5 mL) of each buffer samples and urokinase solution were mixed and incubated for 1 h at 37oC under dark conditions. After incubation, total plasmin activity was measured upon hydrolysis of BODIPY FL-Casein as described above. Data Analysis

All experiments were conducted in quadruplicate on individual milk samples, unless noted above, and analyzed using crossed and nested ANOVA general linear model, with Tukey’s pairwise comparisons at 95% confidence level. Data analysis was performed using Minitab v. 15.2 (Minitab Inc., State College, PA)

RESULTS

Pressure-temperature profiles of milk samples

Figure 2 shows the temperature and pressure history of HPP and PATP milk samples processed at 650 MPa; 32, 78 and 105oC with holding time of 5 min. The initial temperature (Ti) of pressure-treated milk samples was estimated based on previously reported heat of compression values for water at various temperatures, since milk behaves similarly to water under high pressure.16 HPP milk samples

(650 MPa; 32oC for 5 min) had a Ti of 7 ± 1oC and reached the target processing temperature (31 ± 1oC) after 1.5 min of compression. No significant heat loss was observed in these samples as a result of adiabatic compression during pressurization. Similarly, no significant heat loss was observed in HPP milk samples with pressure holding times of 0 and 1 min, respectively (data not shown). Pressure treated samples processed at 650 MPa and 78oC for 5 min were preheated at 48 ± 1oC before pressurization. This temperature allowed milk to reach a target processing temperature of 77 ± 1oC during compression. Unlike HPP milk samples processed at 32oC, pressure treated milk samples processed at 78oC attained higher processing temperatures, possibly due to heat gain experienced during processing. The final processing temperature achieved during pressurization (5 min) for PATP (650 MPa, 105oC) milk samples was 104 ± 1oC.

Proteolysis

Relative proteolysis in raw milk was measured only on 0 d because, as expected, it spoils without any thermal and/or pressure process. Also, the known area of casein bands in raw milk (8.91 ± 0.51 Int*mm2) was not significantly different from that of HTST milk (9.59 ± 0.91 Int*mm2) on 0 d. The extent of relative proteolysis in milk

Fig. 2 Pressure–temperature profiles observed during high pressure treatments of milk. (a) High Pressure Processed (HPP) milk at 650 MPa and 32oC for 5 min. (b) HPP milk at 650 MPa and 78oC for 5 min. (c) And, Pressure-Assisted Thermal Processed (PATP) milk at 650 MPa and 105oC for 5 min. (d) Pressure (MPa). (*)Data shown are means of two independent samples



samples at the end of their shelf life is shown in Figure 3. Proteolysis in pressure-treated samples processed at room temperature (32 ± 1oC) was not significantly different than that of pasteurized milk samples (Figure 3, lanes 2-4). The total area decrease of casein bands of milk samples processed at 650 MPa and 32oC for 0, 1 and 5 min, respectively, ranged from 36.7 to 38.4% at the end of their shelf life (20 d). The area of proteolysis products formed as a result of proteolysis for these samples ranged from 33.4 to 40.7% (Table 2, lines 1-3). These results suggest that pressure treatments at 650 MPa and 32oC are not sufficient to inactivate proteases in milk. Therefore, milk proteins are more likely to undergo changes in proteolysis patterns as a result of enzymatic activity in HPP treated milk.

Pressure-treated samples processed at 650 MPa and 78oC showed the least extent of proteolysis among milk samples (Figure 3, lanes 5-7). Relative proteolysis values for these samples ranged from 31.2 to 33.6% at the end of their shelf life (60 d).

Figure 3 (lanes 8-10) shows the extent of proteolysis in PATP milk. Pressure-treated milk processed at 105oC for 0 min showed the highest proteolysis values among milk samples. Total casein area decrease was 68.9% at the end of 30 days. After this point, these samples were noticeably spoiled and not suitable for further analyses. With increasing holding times, the extent of proteolysis in PATP milk was similar to that of pasteurized and UHT milk. The total casein area decrease in PATP milk samples treated at 105oC for 1 min was 40.06% at the end of their shelf life (45 d). The extent of proteolysis in PATP milk samples treated at 105oC for 5 min was 46.8% on 90 d. Both HPP and PATP treatments yielded milk with proteolysis values similar to those found in our controls. The area of casein bands (as-casein, β-casein, and κ-casein) as percent of total casein

HTST pasteurized milk decreased by 37.2% after 20 d of storage under refrigeration conditions (4 ± 1oC). This value was proportional to the total area increase of proteolysis products formed (36.5%) on 20 d (Table 2, line 10). The total casein area decrease in UHT milk was 34.8% (Figure 3, lane 11). This value was proportional to the area of proteolysis products formed (32.9%) after 90 days of storage at room temperature (Table 2, line 11).

Table 2 Area of proteolysis products formed (Int*mm2) in milk at the beginning (0) and end (f) of storage after treatment

Treatment (Int*mm2)0 (Int*mm2)f

650 MPa, 32oC, 0 min 3.65 ± 0.29 5.13 ± 0.56a

650 MPa, 32oC, 1 min 4.12 ± 0.63 5.50 ± 0.81b

650 MPa, 32oC, 5 min 3.84 ± 0.79 5.33 ± 0.07b

650 MPa, 78oC, 0 min 3.72 ± 0.95 4.99 ± 0.33a

650 MPa, 78oC, 1 min 3.44 ± 0.36 4.60 ± 0.29c

650 MPa, 78oC, 5 min 3.22 ± 0.07 4.37 ± 0.37c

650 MPa, 105oC, 0 min 3.42 ± 0.33 5.66 ± 0.62d

650 MPa, 105oC, 1 min 3.63 ± 0.90 5.08 ± 0.25a

650 MPa, 105oC, 5 min 2.95 ± 0.60 4.28 ± 0.15b

HTST Pasteurized 3.50 ± 0.59 4.77 ± 0.77a

UHT 3.12 ± 0.36 4.14 ± 0.34b

a,b,c,d Average area of proteolysis products formed was measured in triplicate. Different letters in the same line indicate significant differences (P < 0.05).

Fig. 3 Relative percentage decrease of casein bands area as a result of proteolysis in milk at the end of shelf life. Lane 1 is HTST pasteurized milk (77 ± 0.8oC 18 s); lanes 2-4 are High Pressure Processed (HPP) milk (650 MPa, 32oC for 0, 1 and 5 min); lanes 5-7 are HPP milk (650 MPa; 78oC for 0, 1 and 5 min); lanes 8-10 are Pressure-Assisted Thermal Processed (PATP) milk (650 MPa, 105oC for 0, 1 and 5 min); lane 11 is Ultra High Temperature (UHT) milk (138 ± 1oC, 2 s). Different superscripts over the bars indicate significant differences (p≤0.05).



35

in HTST pasteurized milk decreased by 37.2% after 20 d of storage under refrigeration conditions (4 ± 1oC). This value was proportional to the total area increase of proteolysis products formed (36.5%) on 20 d (Table 2, line 10). The total casein area decrease in UHT milk was 34.8% (Figure 3, lane 11). This value was proportional to the area of proteolysis products formed (32.9%) after 90 days of storage at room temperature (Table 2, line 11).

Plasmin activity Figure 4 shows the total concentration of plasmin in milk samples subjected to different pressure-temperature treatments. The concentration of plasmin in raw milk was 0.6 ± 0.03 mg/mL. As mentioned earlier in the introduction section, higher Plasmin activity is observed in milk upon storage as more plasminogen is converted into active Plasmin. Therefore, there was less enzyme activity observed in raw milk samples on 0 d. Pasteurized milk and pressure-treated samples processed at 32oC for 0, 1 and 5 min showed the highest plasmin activity values among milk samples. Total plasmin (plasmin + plasminogen) concentration in pasteurized milk samples was 0.74 ± 0.01 mg/mL; whereas the concentration of the enzyme in pressure-treated samples processed at 32oC ranged from 0.68 to 0.77 mg/mL. Similar plasmin inactivation rates were observed in pressure-treated samples processed at 78oC for 0, 1 and 5 min than in pasteurized milk (p < 0.001). The concentration of plasmin in these samples ranged from 0.66 to 0.68 mg/mL on 0 d.

Plasmin in PATP milk samples showed inactivation levels similar to those found in UHT milk with increasing temperature during the pressure process (Figure 4 lanes 9-12). No significant differences were found between pressure-treated samples processed at 105oC for 0, 1 and 5 min and UHT milk (p < 0.01). The concentration of plasmin in the former samples ranged from 0.53 to 0.56 mg/mL; whereas the concentration of plasmin in UHT milk was 0.54 mg/mL.

In this study, pressure holding time had a significant effect (p < 0.05) on inactivation of the plasmin system in milk. A pressure holding time of 5 min gave the lowest plasmin concentration values (0.53 ± 0.07 mg/mL) among milk samples processed at 650 MPa for 105oC.

DISCUSSION

Proteolysis

Our results are in agreement with previous studies linking the extent of proteolytic activity in milk over time.5 An earlier study showed that 30% of the initial area of casein bands (as-casein, β-casein, and κ-casein) gives rise to formation of proteolysis products in milk after incubation at 37oC for 24 h. The relative percentages of κ-casein did not significantly change over time. However, as-casein and β-casein were rapidly hydrolyzed, possibly into γ-casein residues.17 At the pH of milk (6.6 - 6.7), major proteolytic enzymes, such as plasmin, have optimal activity and they also have remarkable heat resistance characteristics.27 High pressure processing can also enhance proteolysis. Recent studies show that pressure treatments between 100-400 MPa can induce

Fig. 4 Total plasmin concentration in milk immediately after treatments. Lane (1) is raw milk (2% milkfat, homogenized); Lane (2) is pasteurized milk (77 ± 0.8oC , 18 s); Lanes (3-5) are High Pressure Processed (HPP) milk (650 MPa, 32oC for 0, 1 and 5 min); lanes (6-8) are HPP milk (650 MPa; 78oC for 0, 1 and 5 min); lanes (9-11) are Pressure-Assisted Thermal Processed (PATP) milk (650 MPa, 105oC for 0, 1 and 5 min); lane (12) Ultra High Temperature (UHT) milk (138 ± 1oC, 2 s). Different superscripts over the bars indicate significant differences (p≤0.05).



conformational changes in proteins, which increases the amount of proteolysis products formed after treatment.6,23,24 In addition to the pressure applied, the extent of proteolysis by HPP depends on the type of protein. Lactalglobumin appears to be less pressure resistant than lactalbumin during high pressure treatments from 100 to 800 MPa at 20oC. Complete denaturation of lactalglobumin occurs at 600 MPa; whereas lactalbumin is partially denatured (72 %) at 800 MPa for 30 min.11 Our results are in agreement with previous studies reported for HPP treated milk. Pressure treatments at > 250 MPa enhance the disruption of casein micelles in milk.12,23,24,25 The authors attributed denaturation due to structural changes of casein micelles via disruption of hydrophobic and electrostatic interactions under high pressure conditions. However, the extent of proteolysis in milk proteins decreases as a result of increasing processing temperature during HPP treatments.

26,11,12,24 The lesser extent of proteolysis observed in these samples can be attributed to the fact that partially denatured lactalglobumin can form complexes with caseins through disulphide interactions, and therefore obstructs the access of proteases to its substrate.28. At pressures between 400 to 600 MPa, higher rates of enzyme inactivation have been reported with increasing holding time and temperature.29 Our results indicate that pressure treatments (650 MPa) with holding times between 1 - 5 min at 105oC are sufficient to prevent proteolysis in milk to levels close to those found in UHT and pasteurized milk. However, samples with no pressure holding time showed the highest amount of proteolysis products at the end of their shelf life. Storage temperature also influences the extent of plasmin activity in milk. Other studies have reported that plasmin activity is higher in UHT milk samples stored at 30oC than at 20oC after 6 months of storage. The rate of formation of proteolysis products formed was directly

correlated to the storage temperature of the sample.30 In this study, higher proteolysis values were observed in PATP milk samples (650 MPa, 105oC for 0, 1 and 5 min) stored at room temperature (25 ± 1oC) than those of HPP milk samples (650 MPa, 32oC for 0, 1 and 5 min; and 650 MPa, 78oC for 0, 1 and 5 min) stored under refrigeration conditions (4 ± 1oC), as mentioned above.

Plasmin activity

The actual mechanisms of plasmin inactivation under high pressure have not been established yet. The structure of the enzyme under pressure may have a significant role in how proteases are affected by high pressure treatments. Changes in the tertiary and quaternary structure of the enzyme are associated with a reduction in volume during the pressure treatment.31,12 Also, the energy of activation decreases under pressure; and therefore, enzymatic activity is favored as substrate and enzyme are in close proximity to each other. Our observations indicate that higher plasmin concentration values might be the result of a combined inactivation effect of PA’s inhibitors at temperatures close to pasteurization and the pressure treatment applied (650 MPa). Therefore, there was an observed increase of enzymatic activity in milk samples after conversion of plasminogen into active plasmin. Our results are in agreement with previously reported values of plasmin concentration in pasteurized milk. In these studies, the reported plasmin concentration values in milk ranged from 0.14 to 0.73 mg/mL; whereas plasminogen concentration values range from 0.55 to 2.75 mg/mL.4,7 In addition to the considerably high thermal stability of the enzyme, plasmin also shows pressure stability between 100–800 MPa at room temperature conditions and the enzyme shows increased stability at pressures > 600 MPa. The extent of plasmin inactivation



37

under high pressure increases with increasing temperature. The enzyme can be inactivated up to 86.5% after treatments at 400 MPa and 60oC for 15 min.12 Similarly, other authors have reported a synergistic effect of pressure (> 600 MPa) and temperature (85oC) on inactivation of the plasmin system in milk.10 Our results are in accordance with these observations, as PATP milk samples showed higher plasmin inactivation rates compared to HPP milk.

CONCLUSIONS

These results indicate that the pressure–temperature combinations used in this study are sufficient to delay the extent of proteolysis in milk. However, storage conditions have a significant role in the activity of proteases. There is a synergistic effect of pressure and processing temperature on the inactivation of the plasmin system in milk. Although the enzyme was not completely inactivated, combinations of pressure (650 MPa) with higher temperature (105oC) were sufficient to inactivate plasmin to levels similar to traditional UHT processes.

FUNDING AND ACKNOWLEDGEMENTS

The authors would like to thank Smith Dairy Products Co. (Orrville, OH) for providing the milk used in this study.

Author Francisco Parada-Rabell gratefully acknowledges the support given by CONACyT (Mexico) for scholarship # 196181.

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