Greener production of low methoxyl pectin via recyclable enzymatic de-esterification using pectin methylesterase cross-linked enzyme aggregates captured from citrus peels

Journal Pre-proof Greener production of low methoxyl pectin via recyclable enzymatic de-esterification using pectin methylesterase cross-linked enzyme aggregates captured from citrus peels Sachin Talekar, R. Vijayraghavan, Amit Arora, Antonio F. Patti PII:

S0268-005X(19)32184-8

DOI:

https://doi.org/10.1016/j.foodhyd.2020.105786

Reference:

FOOHYD 105786

To appear in:

Food Hydrocolloids

Received Date: 20 September 2019 Revised Date:

15 February 2020

Accepted Date: 17 February 2020

Please cite this article as: Talekar, S., Vijayraghavan, R., Arora, A., Patti, A.F., Greener production of low methoxyl pectin via recyclable enzymatic de-esterification using pectin methylesterase cross-linked enzyme aggregates captured from citrus peels, Food Hydrocolloids (2020), doi: https://doi.org/10.1016/ j.foodhyd.2020.105786. This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. Please note that, during the production process, errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. © 2020 Published by Elsevier Ltd.

Graphical abstract

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Greener production of low methoxyl pectin via recyclable enzymatic de-esterification using

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pectin methylesterase cross-linked enzyme aggregates captured from citrus peels

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Sachin Talekara, b, R. Vijayraghavanc, Amit Aroraa,b*, Antonio F. Pattic* a

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IITB-Monash Research Academy, Indian Institute of Technology Bombay, Powai, Mumbai 400076, India

5 b

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(CTARA), Indian Institute of Technology Bombay, Powai, Mumbai 400076, India

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Bioprocessing Laboratory, Centre for Technology Alternatives for Rural Areas

c

School of Chemistry, Monash University, Wellington Road, Clayton, Victoria 3800, Australia

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*Corresponding authors. Prof. Antonio F. Patti, Prof. Amit Arora

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E-mail address: [email protected], [email protected]

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14 15 16 17 18 19 20 21 22 23

1

1

Abstract

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We propose more green and sustainable enzymatic production of low methoxyl (LM) pectin with

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desired degree of esterification (DE) using a reusable and practically cost-free biocatalyst

4

captured from renewable waste. The pectin methyl esterase (PME) was directly captured as

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recyclable biocatalyst from the waste citrus peels through cross-linked enzyme aggregates

6

(CLEAs). The optimization of biocatalyst preparation specifically recovered 85% of PME

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activity available in the waste citrus peels by ammonium sulphate (50%, v/v) precipitation

8

followed by cross-linking for 5 h with 70 mM glutaraldehyde. The PME-CLEAs were

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characterized by FTIR and SEM and used for batch de-esterification of high methoxyl (HM)

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pectin obtained from citrus, mango, and pomegranate to LM-pectin. The PME-CLEAs exhibited

11

higher stability in acidic pH (most desired for HM-pectin de-esterification), thermostability, and

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similar kinetics of de-esterification compared to the native PME. The PME-CLEAs were

13

extremely versatile as they achieved maximum reduction in DE by 75%, 68%, and 66% for

14

pectin derived from citrus, mango, and pomegranate, respectively without changing their

15

molecular weight (MW) and GalA content under the same optimal batch reaction conditions

16

(pectin loading 1% w/v, PME loading 40 U/g of pectin, 35°C, and pH 6.5, 4 h). De-esterified

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pectin’s strong gelling in presence of Ca2+ and decreased methyl ester peak density in FTIR

18

further confirmed production of LM-pectin by PME-CLEAs. Finally, the PME-CLEAs were

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recycled for 7 batches of LM-pectin production with consistent DE, MW and GalA content of

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LM-pectin produced in each batch and PME activity which makes this process promising to

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pectin industry.

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Key words: Enzymatic pectin de-esterification, Low methoxyl pectin, Pectin methyl esterase,

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Immobilized enzyme, Cross-linked enzyme aggregates

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1. Introduction

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Natural hydrocolloid pectin is widely utilized for thickening, stabilization, and encapsulation in

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food & beverage, cosmetic, and pharmaceutical industries due to which the global pectin market

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is expected to exceed $2.4 billion by 2020 (Ciriminna, Fidalgo, Delisi, Ilharco, & Pagliaro,

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2016). Naturally, the pectin is methyl esterified at the carboxyl group of some of its galacturonic

7

acid residues based on which it is classified into two groups: high methoxyl (HM) pectin (degree

8

of esterification, DE>50%) and low methoxyl (LM) pectin (degree of esterification, DE<50%).

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The gelling of HM-pectin requires a very high concentration of sugar (usually >55%) and highly

10

acidic conditions (pH <3.5) to stabilize the hydrophobic interactions between methoxyl groups.

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On the other hand, in absence of sugar, the LM- pectin gels via the cross linking of free carboxyl

12

groups of galacturonic acids by the metal cations such as Ca2+ in a wide pH range (Chan, Choo,

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Young, & Loh, 2016). Therefore, the demand of LM-pectin for the preparation of low

14

calorie/sugar gels in the dietetic foods and calcium cross-linked LM pectin hydrogels for

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encapsulation and controlled release of drugs, biodegradable and edible food packaging films,

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wound healing, and tissue engineering for hard tissue repair, etc. has increased impressively

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(Espitia, Du, de Jesús Avena-Bustillos, Soares, & McHugh, 2014; Han, et al., 2017; Munarin,

18

Tanzi, & Petrini, 2012).

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The LM-pectin is mainly produced by the chemical de-esterification of HM-pectin with acid,

20

alkali, and alcoholic/aqueous ammonia (Chan, et al., 2016). However, the chemical de-

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esterification suffers from lower reaction rate and selectivity to de-esterification, pectin

22

breakdown, uncontrolled de-esterification making hard to accomplish the desired DE, production

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of LM-pectin with random de-esterification that causes unstable viscosity, phase separation and 3

1

syneresis in aqueous products, chemical remnants in the LM-pectin harmful to consumers,

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production of large amount of chemical waste that damages the environment, and equipment

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corrosion (Chan, et al., 2016; Gerrish, Chambliss, & Forman, 2004). Therefore, an enzymatic de-

4

esterification of HM- pectin by pectin methyl esterase (PME, EC 3.1.1.11) is gaining interest as a

5

green alternative to conventional chemical de-esterification (Cameron, Savary, Hotchkiss, &

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Fishman, 2005; Hua, Yang, Din, Chi, & Yang, 2018; Kim, Williams, Luzio, & Cameron, 2017;

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Wan, Chen, Huang, Liu, & Pan, 2019; Zhao, et al., 2015). Advantages of enzymatic approach

8

include controlled and highly specific removal of methyl esters without the breakdown of pectin

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(Hotchkiss, et al., 2002), block wise de-esterification of HM-pectin that forms stable aqueous

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products (Gerrish, et al., 2004) at mild conditions, and avoiding chemical waste generation

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(Wan, et al., 2019). However, the industrial production of LM-pectin by enzymatic de-

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esterification of HM-pectin still remains a challenge. For successful industrial application, first it

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is necessary to enhance PME productivity by genetic modification and optimization of culture

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conditions (Patidar, Nighojkar, Kumar, & Nighojkar, 2016; Rajulapati & Goyal, 2017; Zhang, et

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al., 2018). Then, expensive and time-consuming purification of PME is essential (Glinka & Liao,

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2011; Plaza, et al., 2007; Zhang, et al., 2018). When the energy consumption, waste generation

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and CO2 emission during the production and purification of PME are also considered, a

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completely different picture evolves questioning the intrinsic greenness of the enzymatic

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production of LM-pectin (Tieves, et al., 2019). Furthermore, the commercial formulations of

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PME are available in soluble form and therefore, not reusable and are employed in a single-use,

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throw-away basis (Cameron, et al., 2005; Hua, et al., 2018; Kim, et al., 2017; Wan, et al., 2019;

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Zhao, et al., 2015), which is not cost-effective (high enzyme cost per kilogram of product) and

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not in line with current philosophy of circular economy and sustainability (Sheldon, 2017). The

4

1

application of immobilized PME can ameliorate the cost-efficacy, since it enables PME recovery

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and reuse (DiCosimo, McAuliffe, Poulose, & Bohlmann, 2013). However, only few attempts

3

have been made for the LM-pectin production using immobilized PME with solid carriers

4

(Sandra Aparecida de Assis, Fernandes, Ferreira, Cabral, & Oliveira, 2004; Sandra Aparecida de

5

Assis, Ferreira, et al., 2004; Nighojkar, Srivastava, & Kumar, 1995) which still suffer from the

6

need of purified PME for the immobilization in addition to the extra cost for carrier, chemical

7

modification and activation of carrier prior to PME immobilization, and activity dilution by large

8

mass (90-99%) of non-catalytic carrier, thus translating to incremented biocatalyst cost,

9

decreased productivity (kg of product per kg of enzyme), and higher environmental footprint

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(Sheldon & Woodley, 2017). For making bio-catalysis sustainable in the food/pharmaceutical

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industries, the method that permits low cost and easy biocatalyst design is truly essential

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(Carceller, et al., 2019). The strategy that can capture naturally available PME directly in

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recyclable form could be an all-inclusive one solution which addresses most, if not all above

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mentioned issues of enzymatic production of LM-pectin.

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Immobilization design of Cross-Linked Enzyme Aggregates (CLEAs) is apparently a prudent

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option owing to the omission of need of purified enzyme and carrier that composes a biocatalyst

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with high productivity. CLEAs with high activity recovery are prepared by selective enzyme

18

precipitation from crude enzyme extract followed by the cross-linking of precipitated enzyme

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that eventually results in one-step purification and immobilization (Cui & Jia, 2015; Sheldon, et

20

al., 2017; Talekar, Joshi, et al., 2013).

21

formulating PME as CLEAs and their potential industrial application for LM-pectin production.

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By meticulous optimization, we recovered the naturally available PME in the citrus waste peels

23

directly into immobilized form of PME-CLEAs, which ultimately eliminates the costs and waste

This motivated us to explore the possibility of

5

1

generation associated with the PME production and purification and making the biocatalyst almost

2

cost-free and bio-catalytic LM-pectin production more green and sustainable. Easy recovery of PME-

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CLEAs by centrifugation/filtration can enable us to stop the reaction at required time and control

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the de-esterification in order to produce LM-pectin with desirable DE. To understand their

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versatility and potential for the industrial production of LM-pectin, the PME-CLEAs were applied for

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the de-esterification of HM-pectin from different sources such as citrus, mango, and pomegranate in a

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batch reactor and their reuse capacity was subsequently studied. It is worth noting that the leftover

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citrus peel residue after the preparation of PME-CLEAs can be further utilized for the extraction of

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HM-pectin which can either be sold as it is or used by PME-CLEAs as a substrate for LM-pectin

10

production. The proposed process can therefore, be easily integrated with the existing industrial pectin

11

production in order to produce the pectin of wide range of DE to meet the market demand (scheme 1).

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2. Materials and methods

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2.1 Materials

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Acetone, acetonitrile, ammonium sulphate, dimethyl sulfoxide (DMSO), ethanol, methanol and

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n-propanol were purchased from Merck. The glutaraldehyde (25%, v/v) and commercial citrus

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pectin (DE 72.4%) were obtained from Sigma Aldrich. The mango pectin (DE 80%) (Banerjee,

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et al., 2018) and pomegranate pectin (DE 62.7%) (Talekar, Patti, Vijayraghavan, & Arora, 2018)

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were obtained by hydrothermal extraction as described previously.

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2.2 Capturing PME from citrus waste peels in CLEAs

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2.2.1 Screening of precipitating agent

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First, the PME was extracted from fresh orange peels following the previous method (Ly-

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Nguyen, et al., 2002) by mixing them (25 g DW) with cold extraction buffer (0.1 M potassium

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phosphate buffer, pH 8 containing 1 M NaCl) in 1:3 (w/v) proportion followed by grinding for 3 6

1

min in laboratory grinder. The resulting slurry was stirred for 12 h at 4°C, centrifuged (9000 rpm

2

for 20 min at 4°C) and filtered to obtain clear crude PME extract. One milliliter of this clear

3

extract contained 20.1 U of PME and 5 mg proteins. Then, the various precipitating agents were

4

first screened for the maximum precipitation of PME from the crude PME extract. The crude

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PME extract (2 mL) was added under continuous shaking with the pre-chilled organic solvents

6

such as acetone, acetonitrile, dimethyl sulfoxide (DMSO), ethanol, methanol, n-propanol and

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saturated ammonium sulphate solution (10 mL each). The mixtures were incubated at 4°C for 1 h

8

for PME precipitation and then centrifuged at 4°C (9000 rpm for 20 min) to collect the

9

precipitate. The precipitates were re-dissolved in 0.5 M potassium phosphate buffer (pH 6.5) to

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measure the activity of PME and protein content. Subsequently, the precipitant which gave

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highest precipitation of PME was selected and its concentration was varied to improve its

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selectivity for PME measured in terms of specific activity and purification. The de-esterification

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of pectin was also determined by treating commercial citrus pectin at 35°C and pH 6.5 for 2 h

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with each precipitate. The treated pectin was precipitated from the reaction mixture by adding

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equal volume of ethanol followed by overnight incubation at 4°C. Then the pectin was collected

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by centrifugation, washed with ethanol, and its DE was determined by the titrimetric method

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described below. For the further study, the precipitating agent showing the highest PME activity,

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specific activity, purification, and reduction in DE of pectin was used.

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2.2.2 Cross-linking of PME

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The saturated ammonium sulphate solution (10 mL each) was added to the crude PME extract (2

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mL) with stirring. The glutaraldehyde was added at various final concentrations (20-100 mM)

22

after 1 h incubation at 4°C and the reaction mixture was incubated for different times (2-6 h) at

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25°C. The PME-CLEAs were separated by centrifugation at 4°C (9000 rpm for 20 min) and the

7

1

supernatants were evaluated for the PME activity. Then, to remove unbound proteins and

2

glutaraldehyde, the CLEAs were washed with 0.5 M potassium phosphate buffer (pH 6.5) and

3

stored at 4°C. The following equation was used to determine the activity recovery of PME in

4

CLEAs:

(%) =













( )

( )

× 100 (1)

5

The pectin de-esterification was determined by treating commercial citrus pectin for 1 h at 35°C

6

and pH 6.5 with PME-CLEAs prepared by different glutaraldehyde concentrations and cross-

7

linking times. Following the centrifugation separation of the PME-CLEAs, the treated pectin was

8

collected from the reaction mixture and its DE was determined using the titrimetric method

9

described below. The percent reduction in DE was calculated based on pectin’s initial DE.

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2.3 Determination of PME activity

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The PME activity was determined titrimetrically by measuring free carboxyl groups produced

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because of the PME action on citrus pectin as described previously (Sandra A de Assis, Lima, &

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de Faria Oliveira, 2001). One unit of PME activity was defined as the amount of enzyme that

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releases 1 µmol of carboxyl groups per minute. The pectinolytic enzyme (polygalacturonase and

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pectin lyase) activity of clear extract was also determined using citrus pectin as a substrate in

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order to check extraction of pectin depolymerizing enzyme, if any. The polygalacturonase

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activity was determined by measuring reducing sugars (equivalent to galacturonic acid) released

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using DNSA method described previously (Deng, et al., 2019). One unit of polygalacturonase

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activity was defined as liberating 1 µmol of reducing sugar per minute at 35°C. Pectin lyase

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activity was estimated by measuring the increase in absorbance of reaction mixture at 235 nm as

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described by Albersheim (1966). One unit of pectinlyase activity was defined as the amount of

8

1

enzyme causing increase of absorbance by 1 unit per minute at 35°C. One milliliter of clear

2

extract contained only 0.4 U of polygalacturonase activity and 0.6 U of pectin lyase activity.

3

4

2.4 Determination of DE of pectin

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The DE of pectin was measured by Food Chemical Codex titration method (Codex, 1996). First,

6

the pectin solution was titrated with NaOH to determine its free carboxylic group titer (V1).

7

Then, the esterified carboxylic group of pectin was de-esterified by NaOH followed by acid

8

neutralization of excess NaOH. Finally, the pectin solution was titrated with NaOH to determine

9

esterified carboxylic group titre (V2). The DE was calculated as the percentage ratio of V2 to the

10

sum of the V1 and V2.

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2.5 Determination of uronic acid content of pectin

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The uronic acid content was determined using spectrophotometric method described previously

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(Talekar, Patti, Vijayraghavan, & Arora, 2019). It was quantified at 525 nm and expressed as D-

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galacturonic acid equivalents in percentage.

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2.6 Determination of molecular weight of pectin

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The average molecular weight of pectin was determined in duplicates by gel permeation

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chromatography (Tosoh High-Performance EcoSEC HLC-8320) according to our previous

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method (Talekar, et al., 2019). Three TSKgel columns (pore size 100 nm, >100 nm, and 10-100

19

nm) connected in series and pullulan standards were used. The experimental conditions included

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solution of 0.1M NaNO3 and 0.1M NaHCO3 as eluent, 1.0 mL/min flow rate, 40°C column

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temperature, and pectin sample 1mg/mL in eluent.

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2.6 Structural characterization of PME-CLEAs

9

1

The molecular structure of PME-CLEAs was analyzed by FTIR. The FTIR spectra of the freez-

2

dried precipitated PME (free enzyme) and PME-CLEAs were recorded on Thermo Scientific

3

Nicolet iS50 FTIR Spectrometer from 4000 to 400 cm-1. The peak frequencies of amide I band

4

within the range of 1600-1700 cm-1 were identified using the second derivative FTIR and the

5

peak areas in amide I were determined by fitting with Gaussian function in Origin 8.0 as

6

previously reported (Pirozzi, Abagnale, Minieri, Pernice, & Aronne, 2016). Lastly, the

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composition of secondary protein structure was measured on the basis of these peak areas. The

8

structural morphology of the PME-CLEAs was studied on JEOL JSM6360 (Germany) scanning

9

electron microscope (SEM) operated at 5 kV. Before scanning, a sputter coater from Emitech

10

K550 (Ashford, UK) was used to coat the dried PME-CLEAs with platinum.

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2.7 Optimal conditions for pectin de-esterification

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The pH influence on the activity of crude PME extract and PME-CLEAs was determined at 35°C

13

in different pH (3.5-8.5) using 0.05 M buffers (pH 3.5-5.5, sodium citrate buffer; pH 6.5-7.5,

14

sodium phosphate buffer; pH 8.5, NaOH/glycine buffer). The effect of temperature on the

15

activity of crude PME extract and PME-CLEAs was determined by varying the temperature of

16

reaction at pH 6.5 from 30-55°C. During this study, the equal amount of activity was used in the

17

form of both crude PME extract and PME-CLEAs for comparison. Finally, the optimum pH and

18

temperature of crude PME extract and PME-CLEAs required for maximum PME activity were

19

determined.

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2.8 Thermal stability, inactivation kinetics and thermodynamics of PME-CLEAs

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The thermal stability of PME was monitored by heating crude PME extract and PME-CLEAs for

22

different times at 30, 40, and 50°C in 0.05 M potassium phosphate buffer pH 6.5 and measuring

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their percent residual PME activity. The percent residual activity of PME at each temperature is

10

1

the PME activity at any time (At) expressed as the percentage of the initial activity (A0). The

2

PME deactivation rate constant (kd) for each temperature was determined as the slope of the plot

3

of natural logarithm of percent residual activity vs. time from the first order enzyme inactivation

4

kinetics equation:

5



6

The half-life (t1/2, time for losing 50% activity) of PME was estimated using equation:

=

%/'



!"#

(2)

= 0.693/,- (3)

7

Furthermore, the thermal stability was also quantified in terms of the deactivation energy (Ed)

8

determined from the slope of the Arrhenius plot: . ,- = .

9

−

-

01

(4)

Where, A = Arrhenius constant, R = gas constant (8.314 J/mol K), T= absolute temperature (K).

10

The change in enthalpy (∆H°), free energy (∆G°) and entropy (∆S°) due to thermal inactivation

11

was determined from Eyring’s transition state theory (Bedade, Muley, & Singhal, 2019): ∆4 =

-

− 01 (5)

8- × ℎ ∆6 = −01. 7 ; (6) 8: × 1 ∆< = =

∆4 − ∆6 > (7) 1

12

Where, KB = Boltzmann constant (1.38 × 10−23 J/K) and h = Planks constant (11.04 × 10−34 J

13

min).

14

2.9 Production of LM pectin in batch mode

15

The HM citrus pectin was dissolved in 0.5 M potassium phosphate buffer (pH 6.5) at different

16

concentrations (0.5-1.25% w/v) to make the 100 mL volume in shake flask. The reaction was

17

performed by adding different PME dosages (20-60 U of PME per gram of HM pectin) in the 11

1

form of both crude PME extract and PME-CLEAs in a thermostatic rotary shaker at 35°C for

2

different times (0.5-4 h). After each reaction time, the resulting pectin was precipitated by adding

3

equal ethanol volume to the reaction mixture, collected from the reaction mixture, and the DE of

4

resultant pectin was measured by titrimetric method. The percent reduction in DE was calculated

5

on the basis of initial DE after each reaction time. In order to check the versatility of PME-

6

CLEAs, the HM pectin extracted from mango and pomegranate waste peels were also used as a

7

substrate for the production of LM pectin in batch mode as described for HM citrus pectin.

8 9 10

2.10 Enzyme kinetics

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PME extract and PME-CLEAs were determined by measuring the initial reaction rate at 35°C

12

with different concentrations of HM pectin (0.01-5 mg/mL, pH 6.5) from different sources such

13

as citrus, mango and pomegranate. The values of Km and Vmax were determined by Graph Pad

14

Prism software from nonlinear regression fitting of the initial reaction rates corresponding to

15

substrate concentrations.

16

2.11 Recycling and reusing of the PME-CLEAs

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To assess the reuse capacity of PME-CLEAs, the PME-CLEAs were employed for the

18

production of LM-pectin from 1% (w/v in 0.05 mM potassium phosphate buffer pH 6.5) HM-

19

pectin from different sources at 35°C. The PME-CLEAs was separated by centrifugation (9000

20

rpm for 20 min) after 4 h of reaction, thoroughly washed with buffer and then applied for the

21

new batch of LM-pectin production. As previously described, the pectin from each batch

22

reaction cycle was collected by ethanol precipitation, its DE was determined by titration, and the

23

percent reduction in DE was calculated on the basis of its initial DE. The reusability of PME-

24

CLEAs was assessed based on the consistency in the DE of pectin (or the percent reduction in

The Michaelis-Menten constant (Km) and the maximum reaction velocity (Vmax) of PME in crude

12

1

DE of pectin) produced in each batch reaction cycle. After each batch reaction cycle, the residual

2

PME activity of the PME-CLEAs was also measured by taking the PME activity of the fresh

3

PME-CLEAs as 100%.

4

2.12 Gelling study

5

Microscopic gelling of original HM pectin and LM pectin produced by PME-CLEAs was tested

6

as described previously (Hua, et al., 2018). Briefly, an appropriate amount of CaCl2 (1M) was

7

added to the 1% (w/v) pectin solution under stirring at 90°C and the set pH value of the mixture

8

was adjusted with NaOH or HCl. Then the pectin solution was transferred into glass sample tube

9

and incubated at 25°C for 2 h. The tube was then inverted to check if there is no flow down of

10

the pectin solution that confirms the formation of microscopic gel.

11

3. Results and discussion

12

3.1 Capturing PME from citrus waste peels in CLEAs

13

To capture the maximum PME activity in CLEAs, a significant amount of PME must be

14

selectively precipitated. We therefore evaluated seven protein precipitating agents such as

15

saturated ammonium sulphate solution, acetone, acetonitrile, dimethyl sulfoxide (DMSO),

16

ethanol, methanol, and n-propanol (10 mL of precipitant added to 2 mL of crude enzyme i.e.

17

83% v/v) for their ability of PME precipitation. The crude enzyme used for precipitation

18

contained 40.2 U of PME together with some pectinolytic activity (0.8 U of polygalacturonase

19

and 1.2 U of pectin lyase). The effectiveness of precipitating agents was also investigated by

20

measuring the reduction in DE of citrus pectin caused by each precipitate. Among the

21

precipitants, the ammonium sulphate enabled full precipitation of the PME activity that showed

22

maximum reduction in DE (Fig. 1a). The lower PME activity of precipitate generated by organic

23

solvents could be due to either the loss of PME flexibility caused by the organic solvent

13

1

penetration (Li, et al., 2018) or the unfolding of PME molecules in organic solvents (Bedade, et

2

al., 2019). However, ammonium sulphate co-precipitated 0.53 U of polygalacturonase (66% of

3

its initial activity) and 0.94 U of pectin lyase (78.3% of its initial activity) along with PME.

4

Capturing pectinolytic activity along with PME in CLEAs is not desirable as it breaks down the

5

pectin. Therefore, the ammonium sulphate concentration was varied to enhance its selectivity for

6

PME. The results showed that 55% v/v ammonium sulphate concentration seems best to

7

precipitate PME selectively and fully with high specific activity and purer form (1.87-fold

8

purification) (Fig. 1b) and at this ammonium sulphate concentration only 0.056 U of

9

polygalacturonase (7% of its initial activity) and 0.1 U of pectin lyase (11% of its initial activity)

10

was precipitated (Fig. 1c). Further increase in concentration of ammonium sulphate beyond 55%

11

v/v showed that precipitation of pectinolytic activities increased and PME precipitation remained

12

almost constant but with a significant decrease in the specific activity and purity. This could be

13

because of shock-wise aggregation helping to retain the three dimensional PME conformation

14

and difference in precipitation of proteins in the crude enzyme extract (Talekar, Joshi, et al.,

15

2013). The ammonium sulfate at 55% v/v concentration was therefore the logical choice as the

16

best precipitating agent for selective capturing of PME in CLEAs.

17

The various concentrations of glutaraldehyde and cross-linking times were tried to achieve the

18

effective cross-linking of PME. As shown in Fig. 2a and 2b, either the glutaraldehyde

19

concentration or the cross-linking time increased the PME activity recovery in CLEAs. The

20

maximum PME activity recovery of 85% was achieved at 70 mM glutaraldehyde concentration

21

and 5 h cross-linking time. While ammonium sulfate at 55% v/v concentration precipitated very

22

less initial pectinolytic activity, we also measured pectinolytic activity of PME-CLEAs obtained

23

at 70 mM glutaraldehyde concentration and 5 h cross-linking time. Interestingly, PME-CLEAs

14

1

prepared under these conditions did not show pectinolytic activity that could be explained either

2

by not having recovered pectinolytic activity due to insufficient cross-linking or inactivation of

3

recovered pectinolytic enzymes due to excessive cross-linking. Figures S1 and S2 show that

4

maximum activity recovery of polygalacturonase (only 6.7%) and pectin lyase (only 10.9%) was

5

obtained in PME-CLEAs after cross-linking with 30 mM and 40 mM glutaraldehyde,

6

respectively, for 4 h. The polygalacturonase and pectin lyase activities were detected in

7

supernatants obtained after cross-linking with glutaraldehyde concentration lower than 30 mM

8

and 40 mM, respectively and cross-linking time lower than 4 h indicating insufficient cross-

9

linking at lower cross-linking conditions. The polygalacturonase and pectin lyase activities

10

significantly decreased in PME-CLEAs and no pectinolytic activity was also detected in

11

supernatants collected after cross-linking conditions exceeding 30 mM glutaraldehyde (for

12

polygalacturonase), 40 mM (for pectin lyase), and 4 h. It could be because of loss of enzyme

13

flexibility and reaction with amino groups required for its pectinolytic activity due to excessive

14

cross-linking at high glutaraldehyde concentrations and cross-linking times (Dal Magro, Hertz,

15

Fernandez-Lafuente, Klein, & Rodrigues, 2016). This indicates that the pectinolytic enzymes

16

could also be recovered in PME-CLEAs obtained by cross-linking with 70 mM glutaraldehyde

17

for 5 h but inactivated due to excessive cross-linking. Thus, although the crude enzyme extract

18

had much less pectinolytic activity the selective precipitation and cross-linking further

19

eliminated the capture of pectinolytic activity in PME-CLEAs. When the glutaraldehyde

20

concentration was exceeded 70 mM or the cross-linking time was exceeded 5 h, the PME-

21

CLEAs activity and the percent reduction in DE of pectin by PME-CLEAs was decreased. This

22

could be due to the restriction of flexibility required by PME for its activity and regidification of

23

PME, which prevents access to the macromolecular substrate such as pectin due to excessive

15

1

cross-linking at high concentrations of glutaraldehyde or longer cross-linking times (Talekar,

2

Pandharbale, et al., 2013). On the other hand, glutaraldehyde concentration and cross-linking

3

time lower than 70 mM and 5 h, respectively, resulted in less PME activity recovery and

4

reduction in DE of pectin by PME-CLEAs which could be attributed to the insufficient cross-

5

linking of precipitated PME (Bedade, et al., 2019). Based on these results, the precipitation using

6

ammonium sulphate at a concentration of 50% v/v followed by cross-linking with 70 mM

7

glutaraldehyde concentration for 5 h was employed for the selective capture of PME in the form

8

of CLEAs for further study. The as-prepared CLEAs were found to have 5.2 U PME activity per

9

milligram.

10

3.2 Structural characterization of PME-CLEAs

11

The appearance of characteristic amid bands of protein (amide I and II) in the FTIR spectra of

12

CLEAs clearly validates the capture of proteins in PME-CLEAs (Fig. 3a). For CLEAs, the slight

13

shift of these amide bands (amide I: 1640 cm-1 shifted to 1650 cm-1 and amide II: 1430 cm-1

14

shifted to 1440 cm-1) reveals the changes in secondary structure of proteins in CLEA. The

15

secondary derivative FTIR analysis of the amide I region (1700-1600 cm-1) in both free PME and

16

PME-CLEAs was used for the secondary molecular structure analysis of proteins. The relative

17

content of secondary structures for free PME and PME-CLEAs was measured on the basis of

18

multi-component peak areas obtained by amide region curve fitting. The PME-CLEAs had lower

19

fraction of α-helix and random structures and higher β-sheets and β-turns content compared with

20

free PME (Table 1). The aggregation/molecular crowding of PME molecules in CLEAs may

21

have resulted in the formation of intermolecular β-sheets. These results imply that the proteins in

22

CLEAs undergo the alteration of secondary structure during the aggregation and cross-linking

16

1

which could lead to the modification of PME activity in CLEAs (Özacar, Mehde, Mehdi, Özacar,

2

& Severgün, 2019; Wang, et al., 2018; Yang, et al., 2019).

3

The morphological structure by scanning electron micrograph of PME-CLEAs showed the

4

formation of spherical structures of the size ranging from 300-500 nm (Fig. 3b). These smooth

5

surface spherical structures are similar to those defined as “type 1 CLEAs” which are like

6

structures in spherical ball (Schoevaart, et al., 2004). The spherical structure of CLEAs exposes

7

maximum enzyme surface, consequently helps to facilitate the accessibility of enzyme’s active

8

site to the substrate. The PME-CLEAs are therefore of type 1 CLEAs that could enable the

9

transfer of pectin to internal enzyme molecules.

10

3.3 Optimal conditions for pectin de-esterification

11

Following the formation of CLEAs, several studies have shown shift in optimum working pH

12

and temperature of enzyme. The PME-CLEAs, however, did not show a change in optimal pH

13

(6.5) and temperature (35°C) over free PME for the pectin de-esterification (Fig. 4). No shift in

14

optimum pH

15

microenvironment surrounding the active site of free PME. Such findings were also observed in

16

case of alcohol and glucose dehydrogenase co-CLEAs (Hu, et al., 2017) and polyvinyl alcohol-

17

degrading enzyme-CLEAs (Bian, et al., 2019). However, the PME-CLEAs showed a broad pH

18

range (> 90% activity retained between pH 5.5 and 7.5) and were also observed to maintain

19

significantly higher activity at acidic conditions (pH < 6.5) than free PME (Fig. 4a). This positive

20

effect of CLEAs on the stability of enzymes under acidic conditions is also observed for

21

combined-CLEAs of pectinolytic enzymes (Goetze et al., 2017). Because the pectin de-

22

esterification will lead to the continuous decline in the pH of reaction mixture, this result allows

23

more efficient application of PME-CLEAs for large scale LM-pectin production in broader pH

indicated

that

the glutaraldehyde cross-linking did

17

not

modify the

1

range compared to the free PME. Similar changes were obtained from the temperature influence

2

study. The results showed that the PME-CLEAs maintained > 80% activity between 35°C and

3

55°C, while the free PME activity decreased to 30% (Fig. 4b). This could be because of the

4

covalent cross-linking of PME molecules in CLEAs that increases their rigidity and protects

5

them from heat exchange denaturation (Dal Magro, Silveira, de Menezes, Benvenutti, Nicolodi,

6

Hertz, Klein, & Rodrigues, 2018).

7

3.4 Thermal stability and deactivation kinetics of PME-CLEAs

8

The PME in free and CLEAs form was exposed separately to 30°C, 40°C and 50°C in 50 mM

9

potassium phosphate buffer pH 6.5 for different times. The temperature dependent activity loss

10

profile is presented in Fig. 5a-c from which the deactivation rate constants (Kd) and half-life (t1/2)

11

of PME were determined (Table 2). The PME-CLEAs were significantly stable at 30-50°C

12

compared to their free counterpart. Specifically, after 24 h, the PME-CLEAs retained 80%, 60%,

13

and 40% of activity at 30°C, 40°C and 50°C, while the free PME retained only 38%, 19.2%, and

14

8.5% activity, respectively. On an average, the PME-CLEAs showed 3.24-fold higher half-life

15

than the free PME in the temperature range of 30-50°C. In addition, as shown in Fig. 5d, the

16

deactivation energy of PME-CLEAs (52.82 KJ/mole) estimated from the slope of the Arrhenius

17

plot in the range of 30-50°C was also higher than that of free PME (37.37 KJ/mole). This

18

enhanced thermal stability of PME-CLEAs could be related to the effective stabilization of the

19

active conformation of PME due to the large number of covalent cross-links between PME

20

molecules in CLEAs, which requires more energy to break down than free PME and prevents

21

denaturation of PME-CLEAs (Fathali, Rezaei, Faramarzi, & Habibi-Rezaei, 2019).

22

The thermodynamic behavior of free PME and PME-CLEAs was also investigated at 30°C, 40°C

23

and 50°C based on the Eyring’s transition state theory. As evident from the decrease in the

18

1

enthalpy (∆H0) with increase in temperature the less energy was required for the inactivation of

2

both biocatalysts at high temperature (Table 2). However, the ∆H0 values of PME-CLEAs were

3

higher than those of free PME, indicating their enhanced thermostability as a result of secondary

4

structural modification of PME in CLEAs form. While ∆H0 represents the enzyme energy barrier

5

to reach transition state of inactivation, the ∆S0 measures the change in disorder of the enzyme

6

structure upon inactivation. The balance between these two parameters determines the

7

spontaneity of the enzyme molecule’s thermal unfolding/inactivation measured in terms of the

8

Gibb’s free energy (∆G0). Overall, high ∆G0 is associated with high resistance to thermal

9

unfolding (de Castro, et al., 2015). The ∆G0 values of the PME-CLEAs were increased by 3.8,

10

3.1, and 2.8% over the free PME at 30°C, 40°C and 50°C, respectively, which further confirms

11

the enhanced thermostability of PME-CLEAs. Apparently, capture of PME in CLEAs generates

12

a better stabilization against heat denaturation compared to free PME. This is of industrial

13

interest as the greater biocatalyst stability will be reflected in the operational stability.

14

3.5 Production of LM-pectin by PME-CLEAs in batch mode

15

The production of LM-pectin in batch mode by de-esterification of HM-pectin using PME-

16

CLEAs and free PME was monitored by measuring the actual DE and percent reduction in DE of

17

HM-pectin derived from waste peels of citrus, mango and pomegranate.

18

The de-esterification at high concentration of HM-pectin is an obvious choice for producing

19

large quantities of LM-pectin. The effect of HM-pectin concentration on its de-esterification was

20

therefore investigated by conducting a series of reactions at 35°C, pH 6.5 by varying the

21

concentration of HM-pectin from 0.5-1.25% (w/v) with 45 U PME/g of HM-pectin (Fig. 6a-c). It

22

was observed that the rate of de-esterification by PME-CLEAs was decreased with an increase in

23

concentration of HM-pectin from all three sources. The free PME also expressed a similar

19

1

decline in de-esterification rate with an increase in concentration of HM-pectin from all three

2

sources. This may be due to the fact that the higher pectin concentrations make the reaction

3

mixture more viscous which could resist the mass transfer of PME-CLEAs and HM-pectin

4

among themselves. There was no significant difference between the rate of de-esterification by

5

free PME and PME-CLEAs up to 1% HM-pectin concentration, but the de-esterification rate for

6

PME-CLEAs was slightly reduced at 1.25% HM-pectin concentration compared to free PME.

7

These results indicated similar mass transfer behavior for free PME and PME-CLEAs up to 1%

8

HM-pectin concentration. In addition, after 4 h reaction with PME-CLEAs, the overall decrease

9

in DE (from 71.42% to 16-18% for citrus, 80% to 22.1-24.7% for mango, and 62.76% to 17.1-

10

19.3% for pomegranate) was similar at 0.5-1% HM-pectin concentration, indicating that

11

although the de-esterification rate is slightly slow at 1%, it works well within a reaction frame of

12

4 h. Consequently, the 1% HM-pectin concentration is an appropriate choice in terms of

13

maximum production of LM-pectin.

14

Although the PME-CLEAs have low cost being derived from the waste and can be recycled, the

15

minimizing PME loading could be further useful for the economic viability of the production of

16

LM-pectin by PME-CLEAs. Fig. 6d reveals the influence of PME loading on the de-

17

esterification of HM-pectin by PME-CLEAs after 2 h reaction. The reduction in DE of HM-

18

pectin was improved from 23.2% to 65.2% for citrus, 19.1% to 49% for pomegranate, and 14.3%

19

to 45% for mango by increasing the PME loading from 20 to 40 U/g HM-pectin. Beyond this no

20

significant improvement of the reduction in DE of HM-pectin was observed. This could be

21

attributed to the saturation of HM-pectin above the PME loading of 40 U/g HM-pectin.

22

The time profile of the decrease in the DE during the de-esterification of 1% HM-pectin derived

23

from citrus, mango and pomegranate waste peels by PME-CLEAs and free PME at the loading

20

1

of 40 U/g HM-pectin is shown in Fig. 7. The de-esterification occurs quickly within the first 2 h,

2

resulting in a reduction in DE of 61-64% for citrus pectin, 50-53% for mango pectin, and 49-

3

53% for pomegranate pectin. The 3 h and 4 h reduction in DE was 74-75% and 75-76% for

4

citrus, 65-66% and 67-68% for mango, and 63-66% and 65-67% for pomegranate. Only slight

5

reduction in DE after 3 h indicates occurrence of the de-esterification reaction equilibrium state

6

due to the less availability of methyl groups for the reaction. However, despite the more

7

availability of methyl groups with mango pectin, the de-esterification reaction rate constant (K)

8

calculated from the integrated rate equation of first order reaction (8 = ln

9

the initial DE and DE is DE at time t) was highest for citrus pectin and similar for mango and

10

pomegranate pectin (Table 3). When we looked at enzyme kinetic parameters of free PME and

11

PME-CLEAs, we found that the affinity (Km) and reaction velocity (Vmax) of both biocatalysts

12

were highest for citrus pectin and similar for mango and pomegranate pectin (Table 4) due to

13

which it may be possible that the de-esterification rate was highest for citrus pectin and similar

14

for mango and pomegranate pectin. However, there was no significant difference between the

15

affinity (Km) and reaction velocity (Vmax) of free PME and PME-CLEAs for pectin from each

16

source, indicating exposure of PME in CLEAs to the substrate without diffusional constrains or

17

reducing its flexibility required to bind to the substrate (Monajati, Borandeh, Hesami, Mansouri,

18

& Tamaddon, 2018). This could be the reason for the similar de-esterification reaction rates of

19

both biocatalysts for each type of pectin at concentration lower that 1% (w/w). Furthermore, the

20

comparison of Km of PME-CLEAs with that of immobilized PME designs from literature in

21

terms of CKm ratios (Km free enzyme/ Km immobilized enzyme (Carceller, et al., 2019)) showed

22

that the CKm value for PME-CLEAs is among the highest reported previously, indicating clearly

23

the advantage of retaining PME affinity by capturing PME in CLEAs form (see Table S1).

%

21

BCD BC

where DE0 is

1

As shown in table 5, apart from the decrease in DE, the de-esterification by PME-CLEAs did not

2

change the galacturonic acid content and molecular weight of pectin derived from citrus, mango,

3

and pomegranate at above optimized batch conditions. This result indicates that PME-CLEAs are

4

able to specifically hydrolyze methyl ester groups and do not depolymerize the pectin during de-

5

esterification, which further confirms no capture of pectinase activity in the PME-CLEAs.

6

3.6 Stability of PME-CLEAs in repeated de-esterification batches

7

The stability of biocatalyst in the repeated use is an important parameter that dictates its

8

economic feasibility. In order to evaluate this stability the PME-CLEAs were used for the

9

production of LM-pectin from HM-pectin of citrus, mango, and pomegranate in repetitive

10

batches at abovementioned optimal conditions. The PME-CLEAs were separated by

11

centrifugation after each batch and used again in the successive LM-pectin production batches.

12

The PME-CLEAs exhibited production of LM-pectin of similar DE from citrus (17.7-18.3%),

13

mango (24.8-25.1%), and pomegranate (19.3-20%) HM-pectin, even after 7 reuse cycles (Fig.

14

8a). Thus, the reduction in DE of HM-pectin from citrus, mango, and pomegranate in each batch

15

was similar. Moreover, the galacturonic acid content (77.8-78.1% for citrus, 70-71.3% for

16

mango, and 74.5-75.7% for pomegranate) and molecular weight (140.1-141.2 kD for citrus, 62-

17

62.7 kD for mango, and 139.2-140.6 kD for pomegranate) of LM-pectin produced in each cycle

18

were similar. It should also be noted that the activity of PME-CLEAs was also stable over the 7

19

reuse cycles. We did not find the leakage of PME activity during each batch reaction. Due to

20

their compression or structural changes, the CLEAs may undergo clump formation during

21

repeated use, hampering their activity in consecutive batches (Talekar, Joshi, et al., 2013).

22

However, we did not observe significant difference between SEM image and FT-IR spectrum of

23

fresh PME-CLEAs (Fig. 3) and PME-CLEAs after 7 cycles of reuse (Fig. 9), indicating that the

22

1

PME-CLEAs were resistant to compression or any structural changes until 7 cycles of repeated

2

use. This excellent stability of PME-CLEAs may be the reason for producing LM-pectin of

3

consistent properties (DE, GalA content, and MW) from citrus, mango, and pomegranate during

4

repetitive de-esterification batches.

5

The longevity and the productivity (mg of HM-pectin de-esterified per mg PME) of the PME-

6

CLEAs are compared with those of different immobilized-PME designs from the literature (see

7

Table S2). As can be seen, PME entrapped in gelatin is not stable and the egg shell adsorbed

8

PME that is stated as stable gives less productivity, indicating that the productivity of our

9

biocatalyst is several times higher than previously reported. This is especially relevant for the

10

industrial LM-pectin production through an eco-friendly and economical method.

11

3.7 Gelling study

12

Usually, LM-pectin gels via the calcium ion cross-linking of free carboxyl groups by following

13

egg-in-box model (Axelos & Thibault, 1991). In order to realize the gel formation in presence of

14

calcium ions, the LM-pectin produced by de-esterification of HM-pectin from different sources

15

using PME-CLEAs (DEcitrus=18.1%, DEmango =25%, and DEpomegranate =19.3%) was used in the

16

microscopic gelling test at pH 3 as described earlier (Hua, et al., 2018). The LM-pectin produced

17

by free PME (DEcitrus=17.8%, DEmango =24.6%, and DEpomegranate =18.8%) was used for

18

comparison. The LM-pectin from different sources produced by both biocatalysts could not gel

19

in absence of calcium ions. The strong microscopic gels were formed at critical calcium ion

20

concentration of 13 mg/g LM-pectin, 22 mg/g LM-pectin, and 17 mg/g LM-pectin produced by

21

PME-CLEAs from citrus, mango, and pomegranate, respectively (Fig. 10). The free PME

22

derived LM-pectin formed strong microscopic gels at critical calcium ion concentration of 11

23

mg/g LM-pectin, 27 mg/g LM-pectin, and 15 mg/g LM-pectin for citrus, mango, and

23

1

pomegranate, respectively. The less calcium requirement for citrus LM-pectin derived by both

2

biocatalysts could be due to its smaller DE. Although the DE of LM-pectin from citrus and

3

pomegranate for both biocatalysts was nearly the same, their calcium ion requirement was

4

different. Furthermore, for the LM-pectin from the same source (mango) the free PME derived

5

LM-pectin required more calcium ions compared to PME-CLEAs derived LM-pectin despite

6

their similar DE. These results could be attributed to the variation in the DE pattern on the

7

backbone chains of LM-pectin derived from different sources and by different forms of

8

biocatalyst (Hua, et al., 2018, Wan, et al., 2019). Thus, these results confirm that the LM-pectin

9

produced by PME-CLEAs can be indeed applied as a gelling agent in dietetic foods in absence of

10

sugar.

11

3.8 FTIR analysis of LM-pectin produced using PME-CLEAs

12

Fig. 11 shows the FT-IR spectra of both LM-pectin produced by PME-CLEAs and native HM-

13

pectin from citrus, mango, and pomegranate. The typical bands at 1000-1100 cm−1, 1740-1745

14

cm−1, and 1630-1632 cm−1 can be attributed to the glycosidic bond C-O stretch, methyl ester, and

15

carboxylic acid groups. The peaks that appear between 3400-3600 cm-1, 2930-2935 cm-1, and

16

1440-1445 cm−1 can be ascribed to the O-H vibration of galacturonic acid hydrogen bonds, C-H

17

and CH2 of methyl group, and CH3 bending of methyl ester, respectively (Hosseini, Khodaiyan,

18

& Yarmand, 2016). There is a significant reduction in the intensity of the peaks at 2930-2935

19

cm-1 and 1740-1745 cm−1 and an increase in the 1630-1632 cm−1 peak’s intensity of LM-pectin

20

produced by PME-CLEAs, which could be related to the removal of methyl ester groups and

21

conversion to acidic groups through de-esterification. Otherwise, the FT-IR spectra of the LM-

22

pectin produced by PME-CLEAs and its native HM-pectin show an excellent correlation with

24

1

each other which indicated that PME-CLEAs specifically catalyze the de-esterification without

2

making other structural changes in the pectin.

3

4. Conclusions

4

For the first time, we captured the in situ PME from biomass waste (citrus peels) directly in the

5

immobilized form “CLEAs” for fully renewable, environmentally and economically sustainable

6

production of LM-pectin via de-esterification of HM-pectin. Our biocatalyst is exceptionally

7

promising given that it is derived from renewable natural waste and its simple preparation which

8

does not require PME production and purification, carrier for immobilization and its

9

functionalization, and extreme pH and temperature, thus overcoming the environmental impact

10

of classical biocatalyst designs and making enzymatic pectin de-esterification greener. Compared

11

to free PME the PME-CLEAs possessed excellent thermostability and tolerance to acidic pH

12

usually generated during production of LM-pectin via de-esterification of HM-pectin. Despite

13

being captured from citrus peels, the PME-CLEAs were exceptionally versatile as they catalyzed

14

the de-esterification of HM-pectin isolated from various sources such as citrus, mango, and

15

pomegranate in batch mode for the production of LM-pectin. The kinetics of LM-pectin

16

production from all three sources demonstrated by PME-CLEAs was similar to the free PME

17

under optimal batch reaction conditions. Easy recovery of PME-CLEAs allows for easy de-

18

esterification control in order to produce LM-pectin of any desired DE. Moreover, the PME-

19

CLEAs were fully active and produced LM-pectin of same DE during 7 consecutive batches of

20

LM-pectin production from each source, thus possessing superior production capacity (kg of

21

LM-pectin per kg of PME) compared to other biocatalyst designs.

22

It is also worth mentioning that due to in situ biocatalyst capture from citrus waste peels which

23

can be further utilized for pectin extraction, the proposed bio-catalytic process of pectin de-

25

1

esterification can be easily integrated into the existing citrus pectin production process in order to

2

produce LM-pectin of wide range of DE. Thus, the proposed bio-catalytic process has great

3

scope for a more economical, sustainable, and greener industrial production of LM-pectin by de-

4

esterification of HM-pectin compared to conventional chemical and enzymatic process.

5

Acknowledgments

6

AA and ST are thankful to the IITB-Monash Research Academy and the Tata Chemicals

7

Innovation Centre, Pune for providing ST with financial support for his PhD study (IMURA

8

0509).

9 10 11 12 13

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33

1

Scheme captions

2

Scheme 1 Schematic illustration of the proposed LM-pectin production by in situ captured PME-

3

CLEAs that can be integrated into existing pectin production industry.

4

Figure captions

5

Fig. 1 (a) Influence of precipitant on precipitated activity of PME and percent reduction in DE

6

obtained by de-esterification of citrus pectin with precipitated PME. Influence of ammonium

7

sulphate concentration on (b) specific activity and purification fold of PME and (c) precipitated

8

pectinolytic activity. The crude extract used for precipitation had 40.2 U of PME which also

9

contained 0.8 U of polygalacturonase and 1.2 U of pectin lyase. The columns/lines (mean values)

10

with different letters are significantly different (p < 0.05).

11

Fig. 2 The activity recovery of PME in CLEAs at different (a) glutaraldehyde concentration and

12

(b) cross-linking time. The right Y axis shows percent reduction in DE obtained by de-

13

esterification of citrus pectin. The 100% activity recovery corresponds to 40.2 U of PME. The

14

columns/lines (mean values) with different letters are significantly different (p < 0.05).

15

Fig. 3 (a) FTIR spectra of free PME and fresh PME-CLEAs and (b) Scanning electron

16

micrograph of fresh PME-CLEAs

17

Fig. 4 Influence of (a) pH and (b) temperature on activity of free PME and PME-CLEAs. In each

18

case the highest PME activity was considered as 100%.

19

Fig. 5 Thermal inactivation of free PME and PME-CLEAs at (a) 30°C (b) 40°C (c) 50°C and (d)

20

Arrhenius plot for free PME and PME-CLEAs.

21

Fig. 6 Batch de-esterification at different concentrations of (a) citrus (b) mango and (c)

22

pomegranate HM-pectin using PME-CLEAs and free PME. Conditions: pH 6.5, 35°C, and PME 34

1

dosage 45 U/g HM-pectin. (d) Effect of dosage of PME-CLEAs on the de-esterification of citrus,

2

mango, and pomegranate HM-pectin. Conditions: pH 6.5, 35°C, and 1% (w/v) HM-pectin

3

concentration. The columns (mean values) for each source with different letters are significantly

4

different (p < 0.05).

5

Fig. 7 Comparative time course of batch de-esterification of (a) citrus (b) mango and (c)

6

pomegranate HM-pectin by free PME and PME-CLEAs. Conditions: pH 6.5, 35°C, 1% (w/v)

7

HM-pectin concentration, and PME dosage 40 U/g HM-pectin.

8

Fig. 8 (a) Recycling of PME-CLEAs for the batch de-esterification of citrus, mango, and

9

pomegranate HM-pectin. Conditions: pH 6.5, 35°C, 4 h, 1% (w/v) HM-pectin concentration, and

10

PME dosage 40 U/g HM-pectin. The columns (mean values) for each source with different

11

letters are significantly different (p < 0.05).

12

Fig. 9 (a) Scanning electron micrograph and (b) FTIR spectra of PME-CLEAs after 7 cycles of

13

reuse.

14

Fig. 10 Microscopic gels of LM-pectin formed via de-esterification of citrus, mango, and

15

pomegranate HM-pectin by PME-CLEAs.

16

Fig. 11 FTIR spectra of native HM-pectin and LM-pectin produced by PME-CLEAs from (a)

17

citrus (b) mango and (c) pomegranate.

35

Table 1. Fractions of secondary structures for free PME and PME-CLEAs. Biocatalyst

Secondary structure α- helix

β- sheets

β- turns

Random structure

Free PME

12.7

33.3

28.6

25.4

PME-CLEAs

8.1

38.4

31.9

21.6

Table 2. Kinetics and thermodynamics parameters of thermal deactivation of free PME and PME-CLEAs. Temperature

Kd (h-1)

t1/2 (h)

Fold increase in t1/2

∆H0 (KJ mol-1)

∆G0 (KJ mol-1)

∆S0 (J mol-1K-1)

Free PME

PMECLEAs

Free PME

PMECLEAs

PMECLEAs

Free PME

PMECLEAs

Free PME

PMECLEAs

Free PME

PMECLEAs

30

0.040

0.010

17.32

69.30

4.00

34.84

50.29

91.43

94.92

-186.6

-147.2

40

0.067

0.022

10.34

31.50

3.04

34.76

50.21

93.18

96.08

-186.5

-146.5

50

0.100

0.037

6.93

18.72

2.70

34.68

50.13

95.17

97.84

-187.2

-147.6

Average of fold increase in the half-life over the range of 30-50°C

3.24

Table 3 Reaction rate constant for de-esterification of HM-pectin from different sources. Rate constant (h-1)

HM-pectin source Free PME

PME-CLEAs

Citrus

0.45 ± 0.03

0.42 ± 0.02

Mango

0.33 ± 0.01

0.31 ± 0.01

Pomegranate

0.32 ± 0.01

0.29 ± 0.02

Table 4 Enzyme kinetics parameters of free PME and PME-CLEAs for HM-pectin from citrus, mango, and pomegranate. Km (mg mL-1)

HM-pectin source

Vmax (µmole min-1)

Free PME

PME-CLEAs

Free PME

PME-CLEAs

Citrus

0.040 ± 0.010

0.047 ± 0.013

11.4 ± 0.75

10 ± 1.1

Mango

0.112 ± 0.027

0.132 ± 0.010

8.3 ± 0.34

7.7 ± 0.79

Pomegranate

0.093 ± 0.011

0.102 ± 0.024

8.8 ± 0.63

8.1 ± 0.47

Table 5 Degree of esterification, GalA content, and molecular weight of parent pectin and LM-pectin produced by PME-CLEAs from different sources. Source

Parent HM-pectin

LM-pectin

DE (%)

GalA (%)

MW (kDa)

DE (%)

GalA (%)

MW (kDa)

Citrus

72.4 ± 2.1

76.3 ± 0.6

141.5 ± 0.6

18.1 ± 1.5

77.9 ± 1.7

140.1 ± 0.8

Mango

80 ± 2.3

67.3 ± 1.8

62.6 ± 0.8

25.1 ± 2.3

69.8 ± 1.3

61.9 ± 0.7

Pomegranate

62.7 ± 1.2

72.7 ± 1.4

140.8 ± 0.7

19.7 ± 1.2

74.2 ± 2.2

139.2 ± 0.9

Scheme 1

Fig. 1a

Fig. 1b

Fig. 1c

Fig. 2a

Fig. 2b

Fig. 3

Fig. 4a

Fig. 4b

Fig. 5a

Fig. 5b

Fig. 5c

Fig. 5d

Fig. 6

Fig. 6d

Fig. 7a

Fig. 7b

Fig. 7c

Fig. 8

Fig. 9

Fig. 10

Fig. 11a

Fig. 11b

Fig. 11c

Highlights

•

Green and sustainable LM-pectin production with reusable and cost-free biocatalyst

•

PME was captured as carrier-free immobilized biocatalyst from waste citrus peels

•

Biocatalyst enhanced PME’s acidic pH and thermal stability without changing kinetics

•

Process produces LM-pectin of customized DE without changing MW and GalA content

•

Biocatalyst was recycled for 7 cycles of LM-pectin production with consistent DE