Permeability of almond cell walls to digestive enzymes

18 Pages·2013·1.12 MB·English

Investigating the permeability of almond
cell walls to digestive enzymes

INFOGEST Short-Term Scientific Missions (STSM) in the Laboratory of Prof.
Frédéric Carrière, the Laboratory of Enzymology at Interfaces and Physiology of Lipolysis,
at the Centre National de la Recherche Scientifique in Marseille

Myriam Grundy
King’s College London

30th September to 1st of November 2013
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Table of contents

Summary of the project..................................................................................... 3
1. Introduction........................................................................................... 4-7
1.1 Almond cells..................................................................................... 4-6
1.2 Oil bodies......................................................................................... 6-7
2. Material and method............................................................................ 7-12
2.1 Separated cell and oil body preparations........................................ 7-8
2.2 Lipolysis of almond lipids.................................................................. 9-11
2.3 Chemical composition of the almond materials…............................ 11-12
3. Results and discussion........................................................................... 12-17
3.1 Identification of endogenous lipase activity..................................... 12
3.2 Lipolysis of almond lipids................................................................... 13-14
3.3 Chemical composition of the almond cell surface............................ 14-17
4. Conclusion............................................................................................. 17
References......................................................................................................... 18

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Summary of the project

Previous studies provide evidence that the physical encapsulation of intracellular nutrients
by intact cell walls of plant foods plays a predominant role in influencing macronutrient
release in human digestion. One unexplored aspect of this is the extent to which digestive
enzymes can pass through the cell-wall barrier.
In this particular project, we will investigate the potential diffusion of various digestive
lipases through almond cell walls and estimate lipid digestion kinetics. To address this
question, the following methodology will be implemented:
-The main lipolytic enzymes found in the GI tract will be tested (gastric lipase,
pancreatic lipase+colipase, pancreatic lipase-related protein 2, carboxyl ester hydrolase,
pancreatic phospholipase A2), alone or in combination to simulate in vitro the gastric and
intestinal phases of digestion
-Kinetic studies to measure the release of lipolysis products from almond cells, as
well as from isolated almond oil bodies used as controls. Lipids will be extracted after
various time of incubation and analysed by TLC and densitometry. The kinetics of almond oil
bodies lipolysis will be also investigated using the pH-stat technique. By comparing the
extent of lipolytic hydrolysis between encapsulated (almond cells) and accessible lipid
bodies (oil bodies), it will be possible to obtain a better insight on the role of cell wall in
nutrient digestion.
-Analysis of variations in the chemical composition of the almond surface by ATR-
FTIR spectroscopy.
Conjointly, microstructural analysis using light and/or confocal microscopy will be
performed to localise the enzymes and verify the integrity of the oil bodies.

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1. Introduction
The exact mechanism of lipase hydrolysis in almond tissue is far from being fully
understood. Hypothetically, the complete digestion of nutrients held within intact cells
relies on the following three steps: diffusion of the enzymes into the plant cells, hydrolysis
of substrate inside cells and diffusion of hydrolysed products out of cells.

1.1 Almond cells
Figure 1 presents the structure of the almond cells within the tissue. The almond cells have
an average diameter of about 35 μm and are surrounded by a cell wall (about 0.1 μm
thickness). The cells are tightly bound together within the cotyledon matrix, when in
solution (e.g. separated) they have a globose conical shape (Figure 2).

Figure 1: Transmission electron microscopy images of almond seed showing oil bodies (white
inclusions), scale bars = 2 µm.

Figure 2: Light microscopy images of separated almond cells
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The major storage protein found in almond (dark inclusions in Figure 1), sometimes called
amandin or almond major protein (AMP), belongs to the legumin (plant casein) class of seed
proteins, itself part of the globulin family. Globulin proteins are classified according to their
sedimentation coefficient, legumin’s being 11S. Amandin accounts for about 70% of the
total soluble proteins. It has a hexameric structure and each of the six subunits is composed
of two polypeptides (α-chain of about 45 kDa and β-chain of about 20 kDa) linked by a
disulphide bridge, giving the molecule a molecular weight of approximately 450 kDa (1).
Lipids, predominantly triacylglycerol, are assembled into oil bodies (OBs, white inclusion in
Figure 1). Depending on the harvest and variety, the kernel is made of approximately 50% of
lipids of which 70-80% is oleic acid, 15% linoleic acid and 5% palmitic acid (2). Compared to
other tree nuts, almond has a low amount of total and saturated fatty acids, but
nonetheless a significant proportion of poly- and monounsaturated fatty acids, with oleic
acid being the predominant fatty acid (3).
Almond seeds carbohydrate (i.e. sugars and starch) and dietary fibre (i.e. cell walls) contents
are about 5.5% and 11.8%, respectively (4). Almond is also rich in micronutrients, mainly
manganese, magnesium, copper, phosphorus and vitamin E. The mineral reserves of the
seed are present in the form of crystals such as calcium oxalate (5).
The cell walls of almond act as a barrier to nutrients release and digestion (4, 6). Plant cells
behave differently when submitted to physical disruption such as mastication, they can
either rupture or separated. Almond cells have been shown to rupture when chewed (4).
However only the outer layers (1 to 3 layers) of the almond particles have their cells
fractures, a high proportion of the cell are still intact when reaching the stomach (Figure 3).

Figure 3: Light (A) and transmission electron (B) microscopy images of the surface of
masticated almond particle.
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Cell walls are made of complex heterogeneous networks of cellulose, hemicelluloses and
pectic substances. The combination of cellulose microfibrils, cross-linking glycan and pectin
network provide strength and rigidity to the cell wall. It is therefore likely that the access of
the substrate (lipids) by lipases will be hindered by the cell wall when the latter is still intact
(Figure 4). It can also be anticipated that the rate and extent of lipid digestion will be
reduced in intact cells compared to lipid readily available (i.e. oil bodies or emulsion).

Figure 4: Schematic representation of the structure of the primary plant cell wall (Davidson,
2005)

1.2 Oil bodies
Almond, similarly to other oilseed plants, store its lipids as TAGs in OBs until they are
eventually mobilised upon seed germination (Figure 5). The TAGs constitute about 50% of
their total dry weight. Oil bodies are small, spherical organelles enclosed in a monolayer of
phospholipids into which unique proteins, mainly oleosins, are embedded (7, 8) (Figure 5B).
Oleosins represent between 1 to 4% of the OB mass; caleosins and steroleosins are also
proteins specific to OBs (9). Oleosins are alkaline proteins of low molecular mass (15 to 26
kDa) that contain three structural domains: (i) an amphipathic N-terminal domain (40-60
residues), (ii) a central hydrophobic domain (72 residues) depicted as anti-parallel β-strands,
and (iii) a C-terminal amphipathic α-helical domain (48 residues) (7). The first domain varies
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significantly between species and it is located at the surface of the oil body. The central
domain is positioned within the TAG matrix at the centre of the oil body whereas the last
domain interacts with the phospholipid layer. Oleosins maintain the integrity of the OBs by
forming a stable amphipathic layer with the phospholipids and thereby prevent coalescence
and aggregation of the OB during desiccation. They also act as a recognition signal for lipase
during germination.

Figure 5: Model of an oil body (A) and the structure of oleosin (B) from corn (7)
Because of the presence of phospholipids on the surface of the oil bodies, the rate and
extent of the lipolysis will be expected to decrease (10). Phospholipids are fundamental
components for the OB stability which is likely to make it more difficult for the gastric and
pancreatic lipase to have access to its substrate (TAG).

2. Materials and methods
2.1 Separated cells and oil body preparations
Separated cells
Almond particles (about 2-3 mm3) were left for 4 weeks to rotate in a solution of 50 mM
cyclohexanediamine tetraacetic acid (CDTA) and a preservative (5 mM sodium metabisulfite,
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Na S O ) at pH 7. The particles were briefly rinsed then mashed using mortar and pestle to a
2 2 5
paste consistency. The sample was loaded on a stack of 3 sieves - apertures 90, 63 and 53
µm and a 20 µm nylon mesh, as well as a base to collect the liquid. After elimination of most
of the water, the material present on the nylon mesh was then transferred into a membrane
(Float-A-Lyzer G2 10 mL, 3.5-5Kd) for dialysis (Figure 6). This process permits the removal of
CDTA from the separated cells as it inhibits lipase activity (11). The membrane was placed in
phosphate buffer (10 mM, pH 7) for 4 to 6 hours, this operation was repeated 4 to 5 times.

Figure 6: Schematic representation of the dialysis process

Oil bodies (OBs)
Oil bodies (OBs) were physically isolated from raw almond seeds using a method previously
described (12). Briefly, almond seeds were homogenised (Laboratory blender, Model
38BL40, Waring Commercial, New Hartford, USA) in water (ratio 1:4) with 1-2 drops of azide
(0.2%) at full power for 2 min. The slurry was filtered through three layers of cheesecloth to
remove almond particles and cell fragments. The filtrate was then centrifuged (Beckman J2-
21 centrifuge; fixed rotor JA-10) at 7500 rpm, 4°C for 20 minutes. The upper layer (creamy
white pad) was removed with a fork and place into bottles/tubes, these were crude oil
bodies.

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2.2 Lipolysis of almond lipids
The enzymes used for the experiments were provided by Prof Carrière, and were as follows:
-Rabbit gastric extract (RGE): specific activity (SA) of 77 U/mg of powder on tributyrin
at pH 5.5, 62 % of lipase per milligram of powder,
-Porcine pancreatic extract (PPE): SA of about 464 U/mg of powder on tributyrin at
pH 8, 5.8 % of lipase per milligram of powder
-Porcine pancreatic lipase (PPL): SA of about 1000 U/mg of powder on tributyrin at
pH 8
-Guinea pig pancreatic lipase-related protein 2 (GPLRP2) has for substrates
galactolipids, monoglycerides, and phospholipids (phospholipase A1 activity). SA of about
160-250 U/mg of powder on galactolipids at pH 8, 1700 U/mg on tributyrin and 500-570
U/mg on mixed micelles PC/bile salts.
The specific activities have been measured on the pH-stat by Dr Sawsan Amara.

Gastrointestinal digestions
Almond materials were added into eppendorf tubes for a weight equivalent to 50 mg of
lipids, which corresponded to about 120 mg of cell preparation and 50 mg of oil bodies.
Each reaction system had a volume of 1 mL as described in Table 1 and were left incubating
for 1h at 37°C. The reactions were performed using RGE and PPE alone, and in combination,
either simultaneously (1h) or one after the other (30 min RGE then 1h PPE), as well as PPL
with colipase.
Quantity Buffer
Concentration
Enzyme of lipase volume pH
(mg/mL)
(μL) (μL)
Gastric phase RGE 1 20 980 5.00
Duodenal phase PPE 20 218 782 6.00
Duodenal phase PPL 20 218 782 6.00
Gastric and duodenal
RGE + PPE as above 20 + 218 762 6.25
phases
Table 1: Experimental conditions for each digestion phase

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Lipid separation
The lipids from the samples were extracted using a chloroform/methanol solution (5:1, v/v),
1 mL aliquot in 5 mL of chloroform/methanol with 200 μL of 1 M HCL. After centrifugation at
2795 g for 10 min, the chloroform layer was collected and 2 spoonful of spatula of MgSO
4
added to absorb any remaining methanol. The tubes were centrifuged for a further 10 min.
The final volume of the organic layer removed was measured and transferred into a 5 mL
vial with a screwcap and kept at−20C until further analysis.
The lipid standards and samples (15 μL) were then spotted on a thin layer silica plate using a
Linomat IV apparatus (Camag, Muttenz, Switzerland) equipped with a 100-μL Hamilton
Syringe. The plate was immerged into a tank containing a mixture of heptane:ether:formic
acid (55:45:1, v/v/v) for neutral lipids, or methyl acetate:1-
propanol:chloroform:methanol:0.25% Potassium chloride (25:25:25:10:7, v/v/v/v/v) for
phospholipids, and left to migrate for about 10 min. Once dried, the plate was sprayed with
a staining solution of copper acetate and phosphoric acid (Cu/H PO 8%). The liquid was left
3 4
to evaporate for 10 min and the plate then placed in the oven at 180°C for 10 min.

Assays of lipase activity
A convenient and well-known method to measure the activity of lipase is the pH-stat
titration (718 Titrino plus Metrohm Ltd., Herisau, Switzerland) (13). The device consists in a
mechanically stirred reaction vessel connected to an electrode that monitors the pH and an
autoburette for the addition of NaOH (Figure 7). The temperature of the reaction system is
maintained constant (37C) via a water bath. The production of FFA following TAG
hydrolysis results in a decrease in the pH of the solution. The amount of NaOH (μmoles)
added as a function of time to keep the pH constant are equivalent to the amount of fatty
acids released due to the lipase activity. If a large excess of substrate is used and if the
enzyme is stable under the selected experimental conditions, linear kinetics of fatty acid
released are observed with time. This activity can be expressed in international units : 1 U =
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