Encapsulation of Pancreatic Lipase in Hydrogel Beads with Self

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Encapsulation of pancreatic lipase in hydrogel beads with self-regulating internal pH microenvironments: Retention of lipase activity after exposure to gastric conditions zipei zhang, fang chen, Ruojie Zhang, Zeyuan Deng, and David Julian McClements J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.6b04644 • Publication Date (Web): 05 Dec 2016 Downloaded from http://pubs.acs.org on December 7, 2016

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Journal of Agricultural and Food Chemistry

Encapsulation of pancreatic lipase in hydrogel beads with self-regulating internal pH microenvironments: Retention of lipase activity after exposure to gastric conditions Zipei Zhang a,1, Fang Chen b,1, Ruojie Zhang a, Zeyuan Deng b, and David Julian McClements a* a

Department of Food Science, University of Massachusetts Amherst, Amherst, MA

01003, USA b

State Key Laboratory of Food Science and Technology, Nanchang University,

Nanchang, No.8 235 Nanjing East Road, Nanchang 330047, Jiangxi, China

1 *

These authors contributed equally to this manuscript. Corresponding author: D.J. McClements (Tel.: +1 413-545-1019; Fax: +1

413-545-1262; E-mail: [email protected]).

1

ACS Paragon Plus Environment


1

ABSTRACT

2

Oral delivery of lipase is important for individuals with exocrine pancreatic

3

insufficiency, however lipase loses activity when exposed to the highly acidic gastric

4

environment. In this study, pancreatic lipase was encapsulated in hydrogel beads

5

fabricated from alginate (gel former), calcium chloride (cross-linker), and magnesium

6

hydroxide (buffer). Fluorescence confocal microscopy imaging was used to map the

7

pH microclimate within the hydrogel beads under simulated gastrointestinal tract

8

(GIT) conditions.

9

moved from mouth (pH 6.3) to stomach (pH < 4), leading to a loss of lipase activity

10

in the small intestine. Conversely, the pH inside buffer-loaded beads remained close

11

to neutral in the mouth (pH 7.33) and stomach (pH 7.39), leading to retention of

12

lipase activity in the small intestine, as shown by pH-stat analysis of lipid digestion.

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The presence of the encapsulated buffer also reduced bead shrinkage under gastric

14

conditions.

The pH within buffer-free beads rapidly decreased when they

15 16

Keywords: alginate; hydrogel beads; lipase; buffer; digestion

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INTRODUCTION Exocrine pancreatic insufficiency (EPI) is a serious condition that accompanies 1, 2

19

several diseases, including chronic pancreatitis and pancreatic cancer

20

when the pancreas does not generate and/or secrete sufficient amounts of digestive

21

enzymes (such as lipases, proteases, and amylases) to appropriately hydrolyze

22

ingested lipids, proteins and carbohydrates, which results in nutritional deficiencies 3.

23

Lipase is one of the most important digestive enzymes derived from the pancreas and

24

it plays a leading role in the digestion of lipids, which typically provide the majority

25

of calories to the human body 4. In addition, lipid digestion and mixed micelle

26

formation in the small intestine is usually an important precursor to the absorption of

27

lipophilic vitamins and nutraceuticals.

28

therapy used to treat individuals suffering from pancreatic insufficiency 5. However,

29

the supplementation of lipase through the oral route is often ineffective because this

30

enzyme is highly susceptible to degradation during passage through the highly acidic

31

environment of the stomach. In healthy individuals, it has been reported that only 1%

32

of lipase activity is retained after passage through the stomach 6. For this reason, there

33

is considerable interest in the development of effective delivery systems that can

34

encapsulate lipase, protect it in the stomach, and then rapidly release it in the small

35

intestine where it can digest any lipids.

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ingested at the same time as a fat-containing meal to increase calorie intake and

37

bioactive absorption.

. EPI occurs

Enzyme supplementation is an important

These delivery systems could then be

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Encapsulation of enzymes within porous polymer matrices have been widely

39

utilized for many years to improve the stability and recovery of enzymes by

40

physically isolating them from the surrounding medium

41

retained within polymer matrices due to physical entrapment or chemical/physical

42

bonding 9, 10. Physical approaches have advantages over chemical approaches because

43

no chemical modification of the polymer of enzyme is required to form a covalent

44

linkage.

45

effectively retain and protect the enzymes without causing pronounced changes in

46

enzyme structure and activity. Hydrogel beads fabricated from biopolymers are

47

particularly suitable for enzyme encapsulation, because they often involve mild

48

preparation conditions to physically entrap or bind enzymes without altering their

49

properties 11. Moreover, hydrogel beads can be prepared from food-grade ingredients

50

(e.g., polysaccharides and/or proteins), which are natural, sustainable, inexpensive,

51

and already acceptable for oral applications

52

encapsulation of bioactive agents can be conveniently fabricated using an

53

extrusion-gelation method

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the bioactive agent (e.g., enzymes) is injected into a cross-linking solution to promote

55

biopolymer gelation.

56

bioactive agents trapped within a biopolymer matrix 15.

7, 8

.

Enzymes may be

Nevertheless, physical approaches still have to be carefully designed to

12

. Hydrogel beads suitable for

13, 14

. In this approach, a biopolymer solution containing

The hydrogel beads formed using this approach consist of

57

A drawback to using conventional hydrogel beads for lipase encapsulation and

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delivery is that the biopolymer matrix is typically highly porous, which allows small 4


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hydrogen ions (H+) to easily diffuse in and out of the beads.

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pH of the beads may quickly become highly acidic within gastric fluids, thereby

61

resulting in acid-induced deactivation of the lipase. Research has been carried out to

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modify the properties of hydrogel beads to improve the pH-stability of encapsulated

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enzymes, such as by crosslinking the hydrogel matrix using specific enzymes or by

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coating the beads with one or more layers of biopolymers through electrostatic

65

deposition

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pH-determining ions (H+ or OH-) through the bead matrix

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approaches are often unsuccessful because the pH-determining ions are still small

68

enough to diffuse through the pores in the hydrogel matrices or coatings, or because

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the release of the enzyme is also strongly inhibited. An alternative approach to

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protecting labile bioactive agents inside hydrogel beads is to use high levels of

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cross-linked proteins (such as caseinates) that can both inhibit the diffusion of H+ ions

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and act as buffering agents 19, 20.

As a result, the internal

16, 17

. These approaches typically attempt to delay the diffusion of 18

. However, these

73

In this study, we fabricated hydrogel beads suitable for lipase encapsulation,

74

protection, and delivery due to their ability to maintain a neutral pH throughout the

75

gastrointestinal tract (GIT).

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beads were created by incorporating an insoluble basic buffer inside them.

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Specifically, the hydrogel beads were fabricated by injecting a mixture of lipase

78

(enzyme), magnesium hydroxide (buffer), and alginate (gelling agent) into a calcium

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chloride (cross-linker) solution under neutral conditions. Mg(OH)2 is insoluble at

These self-regulating pH microenvironment hydrogel

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neutral pH, but dissolves under acid conditions, thereby releasing OH- ions that

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neutralize H+ ions.

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maintain a neutral pH inside the hydrogel beads when they are exposed to gastric

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conditions. Moreover, Mg(OH)2 is regarded as a safe ingredient and is already widely

84

used as an acid neutralizing agent in the food and pharmaceutical industries

85

Alginate was selected as a gelling agent to fabricate the hydrogel beads because it is a

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food-grade ingredient that is already widely utilized for bioactive encapsulation 13, 14.

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The mild cross-linking conditions used to form alginate hydrogels are particularly

88

suitable for the entrapment and encapsulation of enzymes. Furthermore, the

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insolubility of alginate hydrogels under acidic conditions makes them good

90

candidates for the encapsulation and retention of bioactives under acidic gastric

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conditions.

92

Thus, encapsulated insoluble magnesium hydroxide can help

21

.

The potential gastrointestinal fate of lipase loaded into hydrogel beads with and

93

without Mg(OH)2 co-encapsulation was compared. An in vitro gastrointestinal tract

94

(GIT) model was used to simulate mouth, stomach and small intestine conditions. The

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pH inside the hydrogel beads after exposure to different regions within the GIT was

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estimated using a quantitative ratiometric method based on confocal laser scanning

97

microscopy (CLSM). The hydrogel beads developed in this study may provide a new

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strategy for the encapsulation, protection, and delivery of lipase for patients suffering

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from exocrine pancreatic insufficiency.

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MATERIALS AND METHODS

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Materials

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Fluorescein tetramethylrhodamine dextran (FRD, average Mr 70,000) was

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obtained from Molecular Probes (Eugene, OR). Alginic acid (sodium salt), calcium

104

chloride, and magnesium hydroxide were purchased from Sigma Chemical Company

105

(St. Louis, MO). Lipase was purchased from Sigma-Aldrich (Sigma Chemical Co., St.

106

Louis, MO) and as reported by the manufacturer the activity was 100-400 units/mg.

107

Mucin from porcine stomach, porcine bile extract, sodium chloride, monobasic

108

phosphate and dibasic phosphate, Nile red and fluorescein thiocyanate isomer I (FITC)

109

were obtained from either Sigma-Aldrich (Sigma Chemical Co., St. Louis, MO) or

110

Fisher Scientific (Pittsburgh, PA). All chemicals used were analytical grade. Double

111

distilled water was used to make all solutions.

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Hydrogel beads preparation

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An aqueous alginate solution was prepared by dissolving powdered alginic acid

114

(1.6%, w/w) in phosphate buffer and continuously stirring at 60 °C for an hour, then

115

reducing the temperature to 35 °C with continuous stirring until fully dissolved. The

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alginate solution was then mixed with lipase solution to obtain a concentration of 0.8%

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alginate and 2.7% lipase mixture with or without 0.15% Mg(OH)2. For the pH

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mapping, 10 mg/mL FRD in phosphate buffer was added to 0.8% alginate solutions

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(at a volume ratio of 1:200) with or without 0.15% Mg(OH)2. After continuously

120

stirring, the mixtures were injected into 10% calcium chloride solution using a 7



121

commercial encapsulation unit (Encapsulator B-390, BUCHI, Switzerland) with a 120

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µm vibrating nozzle to prepare the hydrogel beads. The encapsulation device was

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operated under fixed conditions: frequency 800 Hz; electrode 800 V; and pressure

124

500 mbar. The formed beads were held in the Ca2+ solution for 30 min at ambient

125

temperature to promote alginate cross-linking and bead hardening.

126

In vitro digestion model

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The lipase-loaded hydrogel beads with or without co-encapsulated buffer (0.15%

128

Mg(OH)2) were passed through a simulated gastrointestinal tract (GIT) that included

129

mouth, stomach and small intestine phases, which was slightly modified from one

130

previously used

131

studied under similar simulated GIT conditions.

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Initial system: 7.5 g of the initial systems were placed into a glass beaker in an

133

incubator shaker at a rotation speed of 100 rpm for 15 min at 37 °C.

134

Mouth phase: 7.5 g of simulated saliva fluid (SSF) containing 0.03 g/mL mucin was

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preheated to 37 °C and then adjusted to pH 6.8. After being mixed with the initial

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samples, the mixture was incubated in an incubator shaker for 2 min at 37 °C to

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mimic agitation in the mouth.

138

Stomach phase: 15 g of simulated gastric fluid was preheated to 37 °C, and then the

139

pH was adjusted to 2.1. After being mixed with 15 g of the sample resulting from the

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mouth phase, the initial pH of the mixture was about 2.5 and incubated in the

141

incubator shaker for 2 h at 37 °C to mimic stomach conditions.

22

. A control sample containing non-encapsulated lipase was also

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Small intestine phase: 30 g of sample resulting from the stomach phase was placed

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into a 100 mL glass beaker that was placed into a water bath at 37 °C and then

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adjusted to pH 7.00. 1.5 g of simulated intestinal fluid was added to the reaction

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vessel, followed by 3.5 g of bile salt solution with constant stirring. The pH of the

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reaction system was adjusted back to 7.00. Then, 2.5 g of 6% (w/w) corn oil-loaded

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emulsions were added to the sample and an automatic titration unit (Metrohm, USA

148

Inc.) was used to monitor the pH and maintain it at pH 7.0 by titrating 0.1 N NaOH

149

solution into the reaction vessel for 2 h at 37 °C. The amount of free fatty acids

150

released was calculated from the titration curves using the following formula:

% FFA = 100 × ( 151

VNaOH × mNaOH × M Lipid WLipid × 2

) ……………………………… (1)

152

Here VNaOH is the volume of sodium hydroxide solution required to neutralize the

153

FFAs produced (mL), mNaOH is the molarity of the sodium hydroxide solution (0.1 N),

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WLipid is the total weight of lipid initially present in the reaction vessel (0.15 g), and

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MLipid is the molecular weight of the corn oil (800 g/mol).

156

Confocal Laser Scanning Microscopy

157

Images were obtained using a confocal scanning laser microscope with a 20 ×

158

objective lens (Nikon D-Eclipse C1 80i, Nikon, Melville, NY, USA). The confocal

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images of the ratiometric dyes used were obtained using 543 and 488 nm excitation

160

wavelengths, and 650 nm/LP and 590 nm/50 emission wavelengths, respectively. All

161

samples were imaged with an exposure time of 0.5 s and a 12.5% excitation power

9



162

level for both channels. The images for each sample were typically acquired in less

163

than 2 min with at least eight measurements per sample. After incubation under small

164

intestine conditions, the oil phase of the samples was dyed with Nile red solution (1

165

mg/mL ethanol). In addition, the lipase was dyed using FITC solution (1 mg/mL

166

dimethyl sulfoxide) prior to measurements by incubating the lipase-loaded beads in

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0.1 mL of FITC dye solution. The excitation and emission spectrum for Nile red were

168

543 nm and 605 nm, respectively and for FITC were 488 nm and 515 nm,

169

respectively. The microstructure images acquired by confocal microscopy were stored

170

and analyzed using image analysis software (NIS- Elements, Nikon, Melville, NY,

171

USA).

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Standard curve preparation

173

A stock FRD solution (10 mg/mL) was prepared by dissolving powdered FRD

174

dye in phosphate buffer (5mM, pH 7) solution. 5 µL/ml FRD stock solution at pH 4–7

175

(adjusted by hydrochloric acid) was imaged using confocal microscopy to determine

176

the dependence of the fluorescence intensity ratio on solution pH. A small aliquot (5

177

µL) of standard solution was added onto a microscope slide for preparation of the

178

standard curve using CLSM. The main advantage of using the CLSM approach is that

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the internal pH of the hydrogel beads can be determined in situ.

180

Image processing for pH measurement

181

The confocal microscopy images were analyzed using Image J software (1.50I,

10


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imagej.nih.gov). The pH inside the hydrogel beads was determined by analyzing the

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average pH inside a single hydrogel bead using repeated scans (at least eight times).

184

The ratio of pixel intensities of two images obtained from two wavelengths (488 nm,

185

543 nm) were calculated and correlated with pH from the obtained standard curve.

186

The images were processed by repeated scans with frame averaging from at least

187

eight measurements.

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Determination of droplet size

189

The particle size distribution of the hydrogel beads with and without

190

co-encapsulated Mg(OH)2 were measured using laser diffraction (Mastersizer 2000,

191

Malvern Instruments Ltd., Malvern, Worcestershire, UK), which is based on analysis

192

of the angular scattering pattern of particulate suspensions. Samples were diluted in

193

aqueous solutions to avoid multiple scattering effects, and then stirred (1200 rpm) to

194

ensure homogeneity. Phosphate buffer (5 mM, pH 7.0) was used to dilute the initial

195

samples, while pH 2.5 adjusted distilled water was used to dilute gastric samples. The

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average particle sizes are reported as the volume-weighted mean diameter (d43).

197

Statistical analysis

198

All experiments were performed on at least three freshly prepared samples. The

199

results are reported as means and standard deviations. Statistical analyses were carried

200

out using Excel (Microsoft, Redmond, VA, USA) and statistical differences (p < 0.05)

201

were established using a statistical software package (SPSS).

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RESULTS AND DISCUSSION

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Development of the standard curve

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In this study, the ability to map the pH values inside the hydrogel beads during

205

their passage through the simulated GIT was necessary so as to understand the

206

physicochemical basis of their ability to retain lipase activity. For this reason, a

207

ratiometric method based on CLSM was developed to quantitatively measure the pH

208

microenvironment

209

Tetramethylrhodamine (FRD) dye was therefore used to estimate the pH change

210

inside the beads. This dye was selected because it has both pH-dependent (FITC) and

211

pH-independent (TMR) fluorescence groups within its molecular structure, which

212

enables the quantification of local pH values based on determination of the

213

fluorescence intensity ratio, after taking into account dye concentration effects

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Furthermore, these two fluorescence groups are conjugated to dextran molecules that

215

have a relatively large molecular weight (around 70,000 Da), which ensures that the

216

dye remains trapped inside the beads. This ratiometric method has previously proved

217

to be successful at measuring the local pH in other types of particles, including pellets

218

and microspheres 25-27.

within

the

hydrogel

beads.

A

fluorescein

and

23, 24

.

219

Preliminary studies of the dye dissolved in buffer solutions confirmed that the

220

emission intensity from the FITC channel decreased as the pH decreased, while the

221

emission intensity from the TMR channel remained constant (data not shown). Over

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the concentration ranges used, the local pH could be obtained independent of dye

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concentration by taking the ratio of the emission intensities at the FITC/TMR

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channels. This is useful since if any of the dye did leak out of the hydrogel beads

225

during fabrication and storage, the TMR/FITC ratio could still be used to determine

226

the pH inside the beads.

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ratio versus pH was established using buffer solutions covering the range pH 4 to 7

228

(Fig. 1). The standard curve clearly shows that the intensity ratio decreases with

229

decreasing pH, and can therefore be used to measure pH in this range.

230

Mapping the pH inside the beads under simulated GIT conditions

In this study, a standard curve of fluorescence intensity

231

An insoluble antacid buffer agent (Mg(OH)2) was encapsulated within the

232

hydrogel beads so as to control the internal pH inside the beads throughout the

233

simulated GIT process. Mg(OH)2 was used because it will be retained as solid

234

particles inside the hydrogel beads under neutral conditions, but will partly dissolve

235

and release hydroxide ions (OH-) when exposed to acid conditions. Hydrogel beads

236

fabricated using similar preparation conditions, but in the absence of Mg(OH)2, were

237

studied as control samples.

238

Mg(OH)2 were then measured after incubation under different simulated GIT

239

conditions (initial, mouth, stomach, and small intestine) using the standard curve

240

described in the previous section (Fig. 1).

241

The pH values inside hydrogel beads with or without

Confocal microscopy images of the hydrogel beads using the FITC and TMR

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channels highlighted changes in their internal pH in different regions of the GIT (Figs.

243

2 and 3).

244

initial samples and the samples after incubation in the mouth phase, irrespective of

245

Mg(OH)2 encapsulation, as shown by the fact that the TMR channel shows that the

246

bead interior had a relatively strong fluorescence intensity. This result can be

247

attributed to the fact that the pH in the initial phase and the mouth phase were fairly

248

similar (pH 7 and 6.8, respectively). After exposure to the stomach phase, the

249

fluorescence intensities of the hydrogel beads containing Mg(OH)2 measured using

250

both the FITC and TMR channels remained relatively strong, which is indicative of a

251

high internal pH.

252

TMR channels were very weak for the hydrogel beads without buffering agent, which

253

suggested that the pH was very low inside the beads.

The pH value inside the hydrogel beads remained fairly constant for the

Conversely, the fluorescence intensities from both the FITC and

254

The average internal pH values of the hydrogel beads calculated from the

255

calibration curve are shown in Table 1. Initially, the internal pH of the hydrogel beads

256

was higher for the beads containing buffer (pH 7.25) than for those without buffer

257

(pH 6.74), which would be expected due to the fact that a small amount of the

258

Mg(OH)2 may have dissolved and released some hydroxyl ions.

259

simulated mouth conditions, the internal pH of the beads remained fairly close to

260

neutral, i.e., pH 6.29 for buffer-free beads and pH 7.33 for buffer-loaded beads. The

261

microscopy ratio images indicated that the fluorescence intensity was uniformly

262

distributed within the buffer-free beads, which indicated that the pH was also 14


After exposure to

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relatively uniform throughout the beads matrix (Fig. 2). Conversely, for the

264

buffer-loaded beads, the fluorescence intensity was more unevenly distributed, which

265

can be attributed to a higher local pH around the insoluble Mg(OH)2 particles (Fig. 3).

266

After incubation in simulated gastric fluids, the fluorescence intensity sharply

267

decreased for the buffer-free hydrogel beads, which suggested that the pH was below

268

the limit of detection (pH < 4) based on the calibration curve (Fig. 1). This result can

269

be attributed to the fact that small hydrogen ions can rapidly diffuse through the pores

270

of the hydrogel beads leading to a low internal pH value. On the contrary, the pH

271

inside the buffer-loaded beads remained relatively high and close to neutral (pH 7.39)

272

after exposure to the simulated gastric fluids. These results suggest that some of the

273

Mg(OH)2 particles slowly dissolved and released hydroxyl ions that were able to

274

neutralize the hydrogen ions arising from the gastric fluids, thereby maintaining the

275

neutral pH conditions inside the beads. The buffer-loaded beads contain about 1.5

276

mg/mL of Mg(OH)2 (i.e., 0.15 w/w%), which corresponds to about 51 mM of OH-

277

inside the beads.

278

submersion of the hydrogel beads in simulated gastric fluids, which suggests that

279

there was sufficient buffer present within the beads to maintain the internal pH over

280

the time foods, supplements, or drugs normally spend in the stomach.

The confocal microscopy images were taken after 2 hours

281

It should be noted that the hydrogel beads remained intact within the mouth,

282

stomach, and small intestine because of the relatively strong cross-links between the

283

alginate molecules and the lack of enzymes to hydrolyze the alginate molecules. 15



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However, the beads should be broken down in the large intestine due to the presence

285

of hydrolytic enzymes released by colonic bacteria that can breakdown dietary fibers

286

such as alginate.

287

Influence of encapsulation on lipase activity

288

In this section, the influence of encapsulation on the ability of lipase to carry out

289

lipid digestion was studied using an automatic titration (“pH-stat”) method.

290

Lipase-loaded beads, with or without co-encapsulated Mg(OH)2, were passed through

291

the mouth and stomach phases, and then they were incubated with emulsified lipids

292

(corn oil-in-water emulsion) under simulated small intestine conditions. The amount

293

of free fatty acids released over time was then calculated from the volume of NaOH

294

added to the samples so as to maintain a constant pH value (7.0).

295

experiment was carried out by exposing free lipase (no hydrogel beads) to the same

296

simulated GIT conditions.

A control

297

For the free lipase and the lipase-loaded buffer-free hydrogel beads, there was

298

almost no free fatty acids released throughout the entire small intestine phase, which

299

suggested that the lipase had been irreversibly deactivated in the stomach phase (Fig.

300

4). The loss of enzymatic activity of lipases upon exposure to low pH conditions has

301

been attributed to the titration of the active site histidine or to the weakening of the

302

coordination of stabilizing calcium ions

303

encapsulation of lipase in alginate beads on its own was insufficient to protect the

28

.

These results suggest that the

16


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304

lipase from acid conditions.

305

fact that small hydroxyl ions (H+) can easily diffuse into beads incubated in simulated

306

gastric fluids, thereby lowering the internal pH and inactivating the lipase, as

307

suggested by the confocal microscopy measurements (Fig. 2, Table 1). The lipid

308

digestion profile of the lipase-loaded beads containing co-encapsulated Mg(OH)2 was

309

appreciably different from the buffer-free beads (Fig. 4).

310

there was a delay in the generation of free fatty acids, which can be attributed to the

311

time required for lipase molecules to diffuse through the beads and into the

312

surrounding medium.

313

release up to about 45 min, after which there was a more gradual increase until a

314

relatively constant final value was attained. These results indicate that the activity of

315

lipase was preserved when it was co-encapsulated with Mg(OH)2 in the hydrogel

316

beads.

317

As mentioned earlier, this effect can be attributed to the

During the first 10 minutes,

After this time, there was a rapid increase in free fatty acids

Further evidence of the ability of the buffer-loaded beads to retain and stabilize

318

lipase was obtained using confocal microscopy (Fig. 5).

319

using FITC, whereas the oil phase was dyed red using Nile red. The confocal

320

microscopy images indicate that most of the lipase was trapped inside the hydrogel

321

beads, i.e., the green (lipase) fluorescence dye was mostly located in the bead interior

322

(Fig. 5). This result suggests that the majority of lipase was successfully delivered to

323

the small intestine phase using the hydrogel beads. Some green fluorescence intensity

324

could also be detected outside the beads in the simulated intestinal fluids, which 17


Lipase was dyed green


Page 18 of 35

325

suggests that some of the lipase was released from the beads.

Presumably, the

326

released lipase was able to adsorb to the surfaces of the emulsified lipids and promote

327

their hydrolysis into free fatty acids.

328

the rate and extent of lipase release from the hydrogel beads under simulated GIT

329

conditions

In future studies it would be useful to quantity

330

Alginate beads have previously been reported to swell when placed in neutral pH

331

solutions because anionic groups on the alginate molecules (carboxylate groups)

332

become highly charged and repel each other causing the hydrogel network to expand

333

29

334

lipase. There were also pronounced differences in the nature of the lipid phase after

335

exposure to small intestine conditions depending on whether the beads contained

336

buffer or not (Fig. 5). For the buffer-free hydrogel beads, the emulsion remained as

337

small individual lipid droplets that surrounded the beads and were difficult to observe

338

in the confocal images because their dimensions were close to the limit of resolution

339

of the microscope.

340

emulsion was converted into large irregular shaped lipid-rich particles, which is

341

characteristic of the mixed micelles formed by lipid digestion

342

suggests that encapsulation of lipase in the buffer-loaded beads maintained its activity

343

throughout the GIT.

.

The relatively large pore size of swollen beads may have promoted the release of

Conversely, for the buffer-loaded hydrogel beads, the initial

18


22

.

This result also

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344

Impact of GIT passage on hydrogel dimensions

345

As mentioned earlier, alginate beads may swell or shrink depending on their pH

346

relative to the pKa value of their charged groups. Calcium alginate tends to shrink

347

under highly acidic conditions because of the reduction in the electrostatic repulsion

348

between the polymer chains when the carboxyl groups become protonated (–COOH,

349

pKa = 3.5). Conversely, they tend to swell at higher pH values because of the

350

electrostatic repulsion between the negatively charged polymer chains. It is therefore

351

interesting to study the impact of external pH on the dimensions of the buffer-free and

352

buffer-loaded hydrogel beads within the simulated GIT.

353

The particle size distribution and mean particle diameter of the hydrogel beads

354

were measured using static light scattering (Figs. 6 and 7).

355

the mean diameter (d43) of the buffer-loaded beads (264 µm) was slightly lower than

356

that of the buffer-free beads (274 µm), which suggested that the presence of the

357

magnesium hydroxide may have caused some shrinkage of the alginate network.

358

Partial dissolution of the buffer may have released some Mg2+ ions that increased the

359

cross-linking of the alginate molecules. Exposure to stomach conditions caused both

360

types of beads to shrink, but the effect was appreciably larger for the buffer-free beads

361

(208 µm) than for the buffer-loaded beads (245 µm).

362

attributed to the fact that the internal pH of the buffer-loaded beads is higher than that

363

of the buffer-free beads, and therefore one would expect the alginate molecules would

364

have a stronger negative charge and greater electrostatic repulsion. 19


For the initial samples,

This difference can be


365

Page 20 of 35

This study has shown that buffer-loaded hydrogel beads (microgels) can be

366

fabricated from food-grade ingredients using simple processing operations.

It was

367

also shown that these beads could be used to improve the stability of encapsulated

368

lipase under simulated gastric conditions, but release it under small intestine

369

conditions, which may be useful for individuals who suffer from exocrine pancreatic

370

insufficiency. Nevertheless, further work is required using animal and human feeding

371

studies to ensure that the buffer-loaded beads can maintain lipase activity under the

372

complex environment of the actual gastrointestinal tract.

373

beads will be subjected to complex shear and compression forces as they pass through

374

a real GIT, which were not mimicked in the current study. In addition, further research

375

is required to ensure that these buffer-loaded beads can be successfully incorporated

376

into supplements or functional food products without reducing their shelf life or

377

quality attributes.

378

ACKNOWLEDGEMENTS

For example, the hydrogel

379

This material was partly based upon work supported by the Cooperative State

380

Research, Extension, Education Service, USDA, Massachusetts Agricultural

381

Experiment Station (MAS00491) and USDA, NRI Grants (2013-03795).

382

REFERENCES

383

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enzyme replacement therapy in chronic pancreatitis. Best Practice & Research

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2. Domínguez‐Muñoz, J. E., Pancreatic exocrine insufficiency: diagnosis and

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treatment. Journal of gastroenterology and hepatology 2011, 26, 12-16.

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3. Guarner, L.; Rodriguez, R.; Guarner, F.; Malagelada, J. R., Fate of oral

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enzymes in pancreatic insufficiency. Gut 1993, 34, 708-712.

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4. Whitcomb, D. C.; Lowe, M. E., Human pancreatic digestive enzymes.

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A.; Vilariño‐Insua, M., Effect of the administration schedule on the therapeutic

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pancreatic insufficiency: a randomized, three‐way crossover study. Alimentary

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pharmacology & therapeutics 2005, 21, 993-1000.

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6. Layer, P.; Go, V. L.; DiMagno, E. P., Fate of pancreatic enzymes during

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7. Zhang, Z.; Zhang, R.; Chen, L.; McClements, D. J., Encapsulation of

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environmental conditions on enzyme activity. Food chemistry 2016, 200,

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13. Giri, T. K.; Thakur, D.; Alexander, A.; Ajazuddin; Badwaik, H.; Tripathi, D.

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to alginate microspheres. Macromolecular bioscience 2005, 5, 717-727.

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17. Taqieddin, E.; Amiji, M., Enzyme immobilization in novel alginate–chitosan

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W.; Chu, L.-Y., Novel intestinal-targeted Ca-alginate-based carrier for

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gelation for the microencapsulation of probiotic cells. International Dairy

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20. Heidebach, T.; Forst, P.; Kulozik, U., Microencapsulation of probiotic cells

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for food applications. Critical reviews in food science and nutrition 2012, 52,

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organic solvents. Journal of biotechnology 2009, 141, 42-46.

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microencapsulation 1996, 13, 319-329. FIGURE CAPTIONS Figure 1. Standard curve of pH vs. intensity ratio taken by 5 µL/ml stock solution (10mg/mL). Figure 2. Fluorescent images of hydrogel beads without buffer encapsulation during the digestion process (intensity of TMR signal was enhanced using Image J to improve contrast). The images of the TMR channel and FITC channel were emitted at 543 and 488 nm, detected at 650/LP and 590/50 nm respectively. Figure 3. Fluorescent images of hydrogel beads with buffer encapsulation during the digestion process (intensity of TMR signal was enhanced using Image J to improve contrast). The images of the TMR channel and FITC channel were emitted at 543 and 488 nm, detected at 650/LP and 590/50 nm respectively.

Figure 4. Amount of free fatty acids released from the systems (free lipase and hydrogel beads with or without buffer co-encapsulation) using a pH-stat in vitro digestion model.

Figure 5. Confocal microscope of lipase-loaded beads with and without buffer encapsulation after exposure to the small intestine phase for 2 hours. Lipase was dyed

25



with FITC to show green fluorescence, while the lipid phase was dyed by Nile Red to show red fluorescence.

Figure 6. Particle size distributions of different samples initially and after the stomach digestion process. Figure 7. Mean particle diameter (d43) of different samples initially and after the stomach digestion process.

26


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Page 27 of 35


Table 1. The Measured pH Value Inside Beads With/ Without Buffer Co-encapsulation During Digestion Process

Initial

Mouth

Stomach

Without buffer (pH)

6.74 ± 0.08

6.29 ± 0.104

pH < 4.0

With buffer (pH)

7.25 ± 0.106

7.33 ± 0.182

7.39 ± 0.15

27



Page 28 of 35

Figure 1

Fluorescence Intensity Ratio

1.6 1.4

Ratio = 9.219 e-0.519pH R² = 0.9781

1.2 1 0.8 0.6 0.4 0.2 0 3.5

4

4.5

5

5.5

6

pH

28


6.5

7

7.5

Page 29 of 35


Figure 2

29



Figure 3

30


Page 30 of 35

Page 31 of 35


Figure 4

90 80

FFA Released (%)

70 60 50 40 Beads (with Buffer)

30

Beads (no Buffer)

20

Free lipase

10 0 0

20

40 60 80 100 Digestion Time (min)

31


120


Figure 5

With buffer

Without buffer

32


Page 32 of 35

Page 33 of 35


Figure 6

70 60 Volume Fraction (%)

initial (no buffer)

50 stomach (no buffer)

40 30

initial (with buffer)

20 10

stomach (with buffer)

0 10

100 1000 Particle Diameter (µm)

33


10000


Figure 7

Mean Particle Diameter (µ µm)

280

b

a

c

260 240

d

220 200 180 160 Initial with buffer

Initial without buffer

Stomach with buffer

34


Stomach without buffer

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Table of Contents Graphic

35


Encapsulation of Pancreatic Lipase in Hydrogel Beads with Self

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