Organocatalytic acetylation of pea starch: Effect of alkanoyl and tartaryl groups on starch acetate performance

Carbohydrate Polymers 294 (2022) 119780

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Carbohydrate Polymers
journal homepage: www.elsevier.com/locate/carbpol

Organocatalytic acetylation of pea starch: Effect of alkanoyl and tartaryl
groups on starch acetate performance
Natalia P. Vidal a, b, Wenqiang Bai a, Mingwei Geng a, c, Mario M. Martinez a, *
a

Center for Innovative Food (CiFOOD), Department of Food Science, Aarhus University, AgroFood Park 48, Aarhus N 8200, Denmark
Aarhus Institute of Advanced Studies (AIAS), Aarhus University, DK-8000 Aarhus, Denmark
c
School of Chemistry and Chemical Engineering, Henan Normal University, Xinxiang 453007, China
b

A R T I C L E I N F O

A B S T R A C T

Keywords:
Organocatalytic esterification
Tartaric acid
NMR
Packaging
Biofilms
Chromatography

Organocatalytic acetylation of pea starch was systematically optimized using tartaric acid as catalyst. The effect

of the degree of substitution with alkanoyl (DSacyl) and tartaryl groups (DStar) on thermal and moisture re­
sistivity, and film-forming properties was investigated. Pea starch with DSacyl from 0.03 to 2.8 was successfully
developed at more efficient reaction rates than acetylated maize starch. Nevertheless, longer reaction time
resulted in granule surface roughness, loss of birefringence, hydrolytic degradation, and a DStar up to 0.5. Solidstate 13C NMR and SEC-MALS-RI suggested that tartaryl groups formed crosslinked di-starch tartrate. Acetylation
increased the hydrophobicity, degradation temperature (by ~17 %), and glass transition temperature (by up to
~38 %) of pea starch. The use of organocatalytically-acetylated pea starch with DSacyl ≤ 0.39 generated starchbased biofilms with higher tensile and water barrier properties. Nevertheless, at higher DS, the incompatibility
between highly acetylated and native pea starches resulted in a heterogenous/microporous structure that
worsened film properties.

1. Introduction
Pea starch is, more than ever before, an abundant by-product from
the increasing production of protein ingredients from field peas, repre­
senting an inexpensive, non-toxic, and annually renewable starch source
compared to wheat, corn, or potato starches (Martinez & Boukid, 2021).
Unfortunately, pea starch demand does not match its escalating abun­
dance due to its inherent properties. As any other starch, pea starch has
several limitations as a replacer of fossil-based polymeric materials,
including its lack of intrinsic thermoplastic behavior (Xu et al., 2020), its
slow recrystallisation after processing that leads to the progressive
embrittlement (Huneault & Li, 2007), and its hydrophilic nature
resulting in poor moisture sensitivity. Furthermore, the inherent water
content of starch can lead to considerable hydrolysis and molar mass
decrease during processing (Imre & Vilaplana, 2020). This deserves
special consideration in those starches with elevated relative proportion
of B-type crystalline polymorphism, such as pea, which possesses 22–55
% of crystals found as B-type allomorph and, hence, with a central cavity
with large amounts of water surrounded by six double helices (P´erez &
Bertoft, 2010; Ren et al., 2021). Pea starch also has the typical limita­
tions of most starches as a food ingredient or drug excipient, such as poor

stability and processing tolerance, high water sorption, low shear, and
heat resistance (Cyras et al., 2006; Parandoosh & Hudson, 1993; Shog­
ren, 1996; Singh et al., 2007), and additional limitations due to its
amylose-driven shortcomings, including low and slow granular swelling
and excessive gel syneresis and stiffness (Martinez & Boukid, 2021).
Interestingly, the three hydroxyl groups in C2, C3 and C6 in the
anhydroglucose units from starch, which confer the hydrophilic nature
to the molecule, are available to be chemically esterified with carboxylic
acids, or carboxylic acid anhydrides or chlorides. Starches having a low
degree of substitution (DS, average number of hydroxyls replaced by
other moieties per repeating unit) find numerous applications in the
food industry as adhesive, thickening, texturizing, stabilizing, and
binding agents (Huang, Schols, Jin, Sulmann, Voragen, 2007a; Imre &
Vilaplana, 2020; Ragavan et al., 2022). Moreover, starch esterified with
short chain fatty acids (e.g., acetate, propionate, butyrate) have the
potential to support the maintenance of a healthy gut (Annison et al.,
2003; Clarke et al., 2011; Nielsen et al., 2019). According to the EU
Regulation (EC) No 1333/2008 (2008), acetylated starch is listed as food
additive (E1420) and can present a maximum level of acetyl groups of
2.5 % as imposed by Commission Regulation (EU) No 231/2012 (2012)
and US Food and Drugs Administration (FDA) (2017), which

* Corresponding author.
E-mail address: (M.M. Martinez).
/>Received 23 February 2022; Received in revised form 20 May 2022; Accepted 22 June 2022
Available online 27 June 2022
0144-8617/© 2022 The Authors. Published by Elsevier Ltd. This is an open access article under the CC BY license ( />

N.P. Vidal et al.

Carbohydrate Polymers 294 (2022) 119780

corresponds to a DS of 0.097. On the other hand, esterified starches with
intermediate (0.2–1.5) and high DS (1.5–3.0) can be used as thermo­
plastic materials with improved thermal stability and reduced moisture
sensitivity compared to native ones (Imre & Vilaplana, 2020).
Currently, commercial starch esters are produced using carboxylic
acid anhydrides and sodium hydroxide (NaOH) as catalyst in aqueous
medium at pH 7–9 (Bello-P´erez et al., 2010; Di Filippo et al., 2016;
Elomaa et al., 2004; Singh et al., 2004; Xu et al., 2004). However, this
starch esterification raises some environmental and safety concerns
since a large volume of wastewater and sodium acetate are generated
(Aˇckar et al., 2015; Ragavan et al., 2022). In this sense, the use of green
organocatalysts with controlled catalytic performance has emerged as a
possible efficient solution to replace NaOH. A wide variety of organo­
catalysts, such as amino acids and hydroxy acids (e.g., lactic, citric, or
tartaric acids), are naturally available from biological sources as single
enantiomers with controlled catalytic performance. This leads to several
remarkable applications in solvent-free and metal-free conditions ideal
to modify biopolymers for food packaging, food, pharmaceutical and
´
biomedical applications (Avila
Ramírez et al., 2019, 2017, 2014). They
can be produced at large scale by biotechnological routes in a straight­
forward and cost-effective manner (Domínguez de María, 2010) and
therefore, are cheap to prepare and readily accessible in a range of
quantities suitable for industrial-scale reactions. Last but not least,
natural organocatalysts are insensitive to oxygen and moisture in the
atmosphere, so there is no need for special reaction equipment and
experimental techniques, and are fully biodegradable, non-toxic and

environmentally friendly (MacMillan, 2008).
Imre and Vilaplana evidenced that, among many organocatalysts,
the hydroxycarboxylic tartaric acid, followed by citric acid, exhibited a
relevant catalytic effect in maize starch esterification (Imre & Vilaplana,
2020). Moreover, tartaric acid catalyst was shown to successfully cata­
lyze the esterification of starch with several 1-substituted mono­
carboxylic acid and anhydride derivatives of n-alkanes, including acetic,
propionic and butyric acids, at DS maximum of 2.93 (Di Filippo et al.,
2016; Tupa et al., 2013, 2015; Nielsen et al., 2018). It must be noted that
in these studies the substrate used was maize starch. Remarkably, the
apparent recalcitrance of native maize starch was greatly influenced by
the role of amylose in stabilizing the semi-crystalline structure of maize
starch and restricting granular swelling (Imre & Vilaplana, 2020; Luo &
Shi, 2012). Although pea starch presents lower granular swelling, less
porous granular structure, higher proportion of B-type crystalline
polymorphism, and considerably smaller amylopectin compared to
normal maize starch (Ren et al., 2021), its recalcitrance towards orga­
nocatalytic derivatization has never been studied.
For the first time, we systematically report the organocatalytic
acylation of pea starch using tartaric acid as catalyst. We hypothesize
that pea starch exhibits lower recalcitrance towards organocatalytic
esterification than maize starch, and that both alkanoyl and tartaryl
substitutions, measured by solid-state 13C NMR, affect the performance
of the resulting starch acetates. Pea starch acetates were studied in terms
of chemical structure, molecular weight, granular morphology, crys­
tallinity, and thermal properties. Moreover, we investigated the role of
the developed tartaric acid-catalyzed pea starches on the mechanical,
thermal and water barrier properties of pea starch-based biofilms.

purity), and lithium bromide were purchased from Sigma Aldrich

(Søborg, Denmark). Hydrochloric acid, analytical grade ethanol, sodium
hydroxide, phenolphthalein, and glycerol were obtained from VWR in­
ternational (Søborg, Denmark).
2.2. Organocatalytic acetylation
The organocatalytic acetylation of pea starch was performed
following other studies focusing on maize starch, such as those from
Imre & Vilaplana and Tupa et al. with some modifications (Imre &
Vilaplana, 2020; Tupa et al., 2015). The pea starch: tartaric acid ratio
was selected from previous studies (Nielsen et al., 2018; Tupa et al.,
2015). L-(+)-tartaric acid (20 g; 0.33 mol) was firstly mixed in 50 mL of
acetic anhydride (0.52 mol) in a 100 mL round flask with a magnetic
stirrer and a reflux condenser to avoid the loss of acetic anhydride. An
initial temperature of 70–80 ◦ C was set using a temperature-controlled
oil bath. When completely dissolved (after 15 min), the temperature
was increased to the desired reaction temperature (85, 95, 110, and
135 ◦ C), and 10 g of freeze-dried pea starch was added. Reaction times
ranging from 30 min to 8 h were tested to evaluate the effect of this
parameter on the degree of substitution of pea starch. After the reaction,
the dispersed mixture was cooled down at room temperature and the
solid material separated from the solvent by vacuum filtration in a
Buchner funnel with Whatman No. 1 filter paper. To ensure the complete
removal of the solvent and organocatalyst, 3 washes with distilled water
and 1 with 50 % ethanol were performed. Washed acetylated starch was
dried overnight in an oven at 40 ◦ C. Dried modified starch was milled
into powder to remove potential aggregates of starch granules. Maize
starch was used as a control to compare with pea starch for the kinetics
of the esterification reaction. Samples were kept under controlled rela­
tive humidity in a desiccator at room temperature until further analysis.
2.3. Determination of the degree of esterification by chemical titration
Back titration with HCl was used to determine the acyl content and

degree of substitution of starch acetates following the procedure from
Tupa et al. (2015). Briefly, 0.11 g of acetylated starch was dispersed in
20 mL of 75 % ethanol and heated at 50 ◦ C for 30 min. pH of the sus­
pensions was decreased with 0.1 N NaOH and phenolphthalein was used
as indicator. Subsequently, 20 mL of 0.1 N NaOH was added prior
subjecting the samples to a second heat at 50 ◦ C for 15 min. Samples
were left under continuous stirring at room temperature for 48 h. After
this time, solutions were back titrated with 0.1 M HCl using the same
indicator. Native pea starch was used as the control. The acyl content
and degree of substitution (DS) were determined as follows:
Acyl (%) =

DS =

(Vc − Vs)*0.1*Macyl *10− 3
× 100
W

(MGlc *Acyl %)
((
)
) × 100
Macyl − 1 *Acyl %
Macyl *100 −

(1)
(2)

Where VC is the volume (mL) of HCl used for the titration of the control,
VS the volume (mL) of HCl used in the sample, Macyl is the molecular

weight of the acetyl groups (43.05 g/ mol), W is the weight of the dried
sample in grams and MGlc is the molecular weight of anhydroglucose
(162.14 g/ mol). Reported DS values were the mean of at least 2
repetitions.

2. Materials and methods
2.1. Materials and reagents
Commercial starch from smooth pea was gently provided by Cosucra
Group Warcoing S.A. (Warcoing, Belgium). Maize starch was purchased
from Ingredion Inc. (Bridgewater, NJ, USA). Pea and maize starch were
freeze-dried for 24 h prior to use to avoid the interference of moisture in
the acetylation process.
Analytical grade acetic anhydride, L-(+)-tartaric acid (>99 % pu­
rity), dimethyl sulfoxide‑d6 (DMSO‑d6), DMSO (HPLC grade, 99.8 %

2.4. Determination of the chemical structure of starch acetates
2.4.1. Fourier Transform Infrared Spectroscopy (FTIR)
Dried starch acetates were analyzed in a Nicolet iS5 Fourier Trans­
form Infrared Spectrophotometer (ThermoScientific, Denmark)
attached to an iD5 Attenuated Total Reflectance (ATR) accessory
(ThermoScienfitic, Denmark). ATR-FTIR was interfaced to a personal
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Carbohydrate Polymers 294 (2022) 119780

computer operating under OMNIC 9 software (version 2.11). FTIR
spectra of native and esterified starch were acquired between 400 and

4000 cm− 1 at a resolution of 4 cm− 1 using 32 co-added scans. The
assignment of the bands to the specific functional group vibration mode
was made by a comparison to previous studies (Tupa et al., 2013; 2015).

2.6. X-ray powder diffraction patterns
The powder X-ray diffraction pattern of the samples were analyzed
using a Bruker D8 Discover A25 diffractometer (Bruker AXS, Rheinfel­
den, Germany) equipped with a copper tube operating at 40 kV and 30
mA, producing CuKa radiation of 0.154-nm wavelength. The diffracto­
grams were collected in a 2θ angle ranging from 5 to 40◦ with a step size
of 0.02◦ .

2.4.2. Nuclear magnetic resonance
2.4.2.1. Proton Nuclear Magnetic Resonance (1H NMR). Native and
acetylated starch (10 mg) were dissolved in deuterated dimethyl sulf­
oxide (DMSO‑d6, 1 mL) (Sigma-Aldrich A/S, Copenhagen, Denmark),
and incubated at 100 ◦ C until a complete solubilization was achieved.
Immediately before analysis, 600 μL of clear solutions were transferred
to 5 mm NMR tubes. Samples were analyzed on a Bruker Avance III NMR
operating at 600.13 MHz. The experimental conditions were spectral
width, 5000 Hz; relaxation delay, 5 s; number of scans, 64; pulse width,
90◦ , with a total acquisition time of 8 min and 49 s. The experiments
were carried out at 25 ◦ C. Due to the hygroscopic nature of the
DMSO‑d6, and to avoid starch retrogradation, samples were freshly
prepared before each analysis. Residual non-deuterated DMSO signal at
2.549 ppm was used as a reference. The spectra obtained were analyzed
using MestReNova software (version 14.2.1) (Mestrelab Research S.L.,
Santiago de Compostela, Spain). The acetylation degree of pea starch
acetates was calculated by using the ratio of the signal area of the pro­
tons corresponding to the -CH3 acyl group (Aacyl) centered at 2.07 ppm

and the area of the signals between 3.2 and 5.5 ppm representing the
protons of anhydroglucose units (AGlc), by using the following equation:
/
Aacyl 7 3*Aacyl
DS =
(3)
=
7*AGlc
AGlc /3

2.7. Molecular size distribution and weight average molecular weight
(Mw)
Molecular weight distribution of fully branched native and acety­
lated starch was determined following the procedures described by
(Martinez et al., 2018) and (Roman et al., 2019). Briefly, 8.0 ± 0.5 mg of
starch was dissolved in 1.5 mL DMSO (Sigma, 99.8 % purity) at 80 ◦ C in
a thermomixer (Eppendorf, Hamburg, Germany) at 350 rpm for 24 h.
Samples were then centrifuged at 4800 rpm for 15 min and the super­
natant was collected. Starch was precipitated with 10 mL 95 % ethanol,
the pellet was collected after centrifugation (4800 rpm, 15 min, 4 ◦ C)
and resuspended in 1.5 mL of DMSO containing 0.5 % lithium bromide
(w/w). The mixture was dissolved at 80 ◦ C and 350 rpm overnight. After
centrifugation (7000 rpm, 10 min), the supernatant was transferred to a
vial for further analysis by High Performance Size Exclusion Chroma­
tography (HPSEC, Agilent 1260 Infinity II, Agilent Technologies,
Waldbronn, Germany) connected to a Multi-Angle Laser-Light Scat­
tering detector (MALS) (Wyatt Technology, Santa Barbara, CA) and a
refractive-index (RI) detector (Shodex RI-501, Munich, Germany). An
injection volume of 100 μL of starch solution was eluted using DMSO/
LiBr as solvent at a 0.3 mL/min flow rate and 80 ◦ C. Size separation was

performed in GRAM 3000 and GRAM 30 (PSS GmbH, Mainz, Germany)
columns connected in series. Data to calculate weight average molecular
weight (Mw) were analyzed by ASTRA software (version 8.1; Wyatt
Technology, Santa Barbara, CA) using a second-order Berry plot pro­
cedure. The specific refractive index increment (dn/dc) was set as 0.066
mL/g as previously reported in other starch dissolved in DMSO (Roman
et al., 2019) and second viral coefficient (A2) was assumed to be
negligible.

Aacyl and AGlc were normalized according to the number of protons (3
and 7, respectively) contributing to the area of the spectral signals.
2.4.2.2. Solid state SP and CP/MAS 13C magnetic resonance. Native and
acetylated pea starch were analyzed by solid-state 13C NMR following
the procedure from (Nielsen et al., 2018). All solid-state NMR spectro­
scopic experiments were performed in a Bruker Avance 400 NMR
spectrometer operating at 400.13 MHz and 100.63 MHz for 1H and 13C
respectively. Solid State Single Pulse (SP) and Cross-Polarization Magic
Angle Spinning (CP/MAS) NMR experiments were recorded at 300 K. A
CP/MAS probe for 4 mm rotos using a spin-rate of 9 kHz, a radio fre­
quency of 70 kHz, spectral width of 50.13 kHz was selected. 1H
decoupling was applied. Recycle delays of 16 and 128 s were used for CP
and SP/MAS, respectively. Determination of the degree of substitution
of starch acetates (DSacyl) and the formation of starch tartrates (DStar)
was done by calculating the ratio of the areas of the signal due to the
carbon of the acyl group bonded to the carbonyl group (16.56 ppm) or to
the carbonyl carbon in the acid/ester (166.88 ppm), respectively, to that
of the glucose anomeric carbon of the anhydroglucose units in the starch
(91.02 ppm). The ratios were determined from the spectra of 13C SP/
MAS NMR due to the different longitudinal relaxation times of the
hydrogen bearing carbons and the carbonyl groups (Nielsen et al.,

2018).

2.8. Thermal properties
Thermal properties of native and acetylated starch samples in dry
state (moisture content <2 % in all the samples) were investigated by
differential scanning calorimetry (DSC) and thermogravimetric analysis
(TGA). DSC thermograms were obtained in nitrogen using a DSC Q2000
(Thermal Advantage Instruments-Waters LLC, USA). Dried starch (6 mg)
was placed in an aluminum pan and hermetically sealed. All the samples
were subjected to heating/cooling cycles in a temperature range of
30–300 ◦ C at heating and cooling rates of 10 ◦ C/min and 50 ◦ C/min,
respectively. Glass transition temperature (Tg) was determined in the
second heating run to eliminate any thermal history in the samples.
Thermal stability in the native and acetylated starch was evaluated using
a thermogravimetrical analysis system TGA-2 STARe from Mettler
Toledo (USA) equipped with STARe software. 5–6 mg of sample were
placed in Mettler Toledo aluminum crucibles with punctured lids and
heated at a rate of 10 ◦ C/min from 25 ◦ C to 600 ◦ C under a high-purity
nitrogen environment. The first derivative analysis (DTG) was per­
formed and the peak temperature at which the thermal decomposition
occurred was determined.

2.5. Microstructure and birefringence
A small amount of pea starch samples was placed on a glass micro­
scope slide with a drop of water and covered by a cover slip. Crystalline
and granular structure, as well as birefringence, of the native and
modified starches were observed by an optical microscope (Leica
Microsystems, Wetzlar, Germany) linked to an Infinity 3 camera and
controlled by the Infinity Analyze 6 software (version 6.0.0., Teledyne
Lumenera, Ontario, Canada). Images were captured with and without

polarized light with a 20× objective.

2.9. Biofilm-making and properties
Starch-based biofilms were developed by the solvent casting method
as reported elsewhere (Saberi et al., 2016). Starch acetates produced at
different reaction times (from 0.5 to 4 h), and resulting in DSacyl values
ranging from 0.39 to 2.80 (Table 1), were used in combination with
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Carbohydrate Polymers 294 (2022) 119780

Table 1
Weight average molecular weight (Mw) and degree of substitution of acyl (DSacyl) and tartaryl groups (DStar) at 135 ◦ C calculated by back titration, 1H NMR, and SinglePulse 13C NMR (SP 13C NMR).
Titration

1

Time (h)

DStotal

DSacyl

DSacyl

DStar

DStotal

Starch acetate (%)

Starch tartrate (%)

Mw (105 g/mol)

0
0.5
1
2
3
4

–
0.39 ±
1.26 ±
2.83 ±
3.16 ±
3.13 ±

–
0.22
0.77
2.04
2.38
2.30

–

0.39
1.00
2.23
2.80
2.63

–
0.0
0.05
0.35
0.39
0.47

–
0.39
1.05
2.58
3.19
3.11

100.0
100.0
90.9
84.6
90.3
83.8

–
–
9.1

15.4
9.7
16.2

74.9 ±
2.65 ±
0.78 ±
0.63 ±
0.57 ±
3.50 ±

0.00
0.07
0.07
0.30
0.54

H NMR

SP

13

C NMR

SEC-MALS
3.13
0.50
0.11
0.31

0.54
2.13

*DSacyl and DStar were determined as described in Section 2.4.2. Starch tartrate DS (DStar) was calculated by the difference between DStotal and DSacyl values.

native starch (1:3 acetylated:native starch ratio, w/w) due to the poor
solubility of highly acetylated starch. Briefly, 4.0 g of starch (or the
native:acetylated starch blend) was mixed with 100 mL of deionized
water containing 25 % of glycerol (w/w of total solids) as the plasticizer.
The film-forming solution was prepared by incubating the mixture at
80 ◦ C for 15 mins under vigorous stirring. To ensure a complete gela­
tinization of the starch and the good homogenization of the solution, the
film-forming solution was heated in a shaking water bath at 95 ◦ C for 1 h
(JULABO, SW22, Germany).
After incubation, the solution was cooled down at room temperature
for 30 min under gentle stirring and degassed using an ultrasonic bath
for 10 min. Then, 40 mL of film-forming solution was gently poured into
bottom-flat 14 cm diameter plastic petri dishes, and oven-dried at 35 ◦ C
for 15 h under circulating air. After this time, biofilms were manually
peeled off from the petri dishes and kept in a desiccator with saturated
K2CO3 salt solution (Relative Humidity = 43 %) at room temperature
until further analysis.

2.10. Statistical analysis
Statistical analysis was conducted using XLSTAT premium software
(version 2021.1.1, New York, USA). Differences among the starch and
biofilm samples parameters were studied by analysis of variance
(ANOVA). When significant differences (p < 0.05) were found, Fisher's
Least Significant Difference (LSD) posthoc test was used to assess the
differences in the means at a significance level of 5 % (α = 0.05). Re­

lationships between degree of substitution in acetylated pea starch and
starch or biofilm properties were performed by Pearson's correlation.
3. Results and discussion
3.1. Optimization of the tartaric acid catalyzed acetylation process
With a view to synthesizing tartaric acid-catalyzed acetylated pea
starch with different degree of substitution (DS), reaction temperature
and time were systematically tested for DS values measured by the
classical titration method. The type of reagent (i.e., acetic acid anhy­
dride) and catalyst (i.e., tartaric acid) were selected based on their high
reactivity and catalytic efficiency, respectively, as shown in previous
studies (Imre & Vilaplana, 2020). The temperature ranged from 85 to
135 ◦ C with a reaction time up to 8 h. The DS increased as a function of
these two factors, as reported by other studies (Di Filippo et al., 2016;
Imre & Vilaplana, 2020; Tupa et al., 2013). However, temperature
showed a higher acylation effect than reaction time. More specifically, a
temperature of 85 ◦ C during 3, 4, and 6 h of reaction led to DS of 0.03,
0.05 and 0.09, respectively. An increase in the temperature to 95 and
110 ◦ C resulted in an increase of the DS values to 0.13 and 0.28, which
were further increased with a longer reaction time (8 h) to 0.41 and
0.49, respectively (see Fig. 1). Results therefore showed that acetylated
starches with low (≤0.2) and even intermediate DS (0.2–1.5) can be
attained at temperatures ≤110 ◦ C. Presumably, esterification of weakly
substituted (DS < 0.2) granules only occurs in the amorphous parts of
the amylopectin fraction exclusively present in the outer lamella of the
granules, as reported elsewhere for base-catalyzed smooth pea acety­
lated starch at low DS (Biliaderis, 2002; Huang, Schols, Jin, Sulmann,
Voragen, 2007b).
To achieve higher substitution levels, the reaction temperature was
increased to 135 ◦ C, temperature just below the boiling point of acetic
anhydride (139.5 ◦ C), and DS values were investigated at reaction times

ranging from 30 min to 4 h. The reactivity significantly raised, reaching
0.39 DS in the first 30 min. Furthermore, DS dramatically increased up
to 2.83 after 2 h of reaction, with a minimal improvement achieved after
3 h. These findings align with those from Tupa et al., who attained DS of
2.93 at 130 ◦ C for 3 h using tartaric acid as catalysts and maize as the
starch source (Tupa et al., 2015). Counterintuitively, maize starch was
more recalcitrant than pea starch (Fig. 1), despite the well-known pores
and channels of the former, and the lack of porosity of the latter (Ren
et al., 2021). These results agree with other studies showing significantly
lower DS (1.23) of maize starch derivatized at 120 ◦ C for up to 7 h (Tupa
et al., 2013). Likewise, Imre & Vilaplana reported maximum DS of 2

2.9.1. Biofilm thickness and mechanical properties
A portable digital dial pipe gauge (Diesella, Denmark) with the ac­
curacy 0.01 mm was used to measure the thickness of the films. Eight
measurements were randomly taken at different points on each film
specimen. The mechanical properties of the native and acetylated
starch-based biofilms were tested at room temperature in a Texture
Analyzer (Mecmesin, FTC, USA) with a 500 N load cell following ASTM
D882 with slight modifications (ASTM 882-01, 2001). Films were cut in
rectangular strips of 80 × 20 mm. The strip was clamped between filmextension grips (MECMESIN 500 N wedge grips, KYOCERA UNIMERCO
Tooling A/S, Denmark) which were set 50 mm apart. The stretching
speed was 10 mm/min. Force-distance curves were obtained and
transformed into stress-strain curves which allowed tensile strength at
break (TS, MPa), percentage of elongation at break (EB, %) and elastic
modulus (EM, MPa) to be obtained. Mechanical properties were calcu­
lated as the average of eight repetitions.
2.9.2. Thermal stability and gas barrier properties
Thermal stability of the biofilms was evaluated using a thermogra­
vimetrical analysis system TGA-2 STARe from Mettler Toledo (USA)

equipped with STARe software, as described in Section 2.8 for pure
starches. The water vapor permeability (WVP) of the film was deter­
mined by a permeability analyzer linked to a pressure-modulated
infrared detector (Totalperm, Permtech Srl, Italy) according to ASTM
F1249-13 (2013) (23 ◦ C and 85 % relative humidity, RH) following
Ellingford et al. methodology with some modifications (Ellingford et al.,
2022). Briefly, before each measurement, the two semi-chambers were
initially preconditioned with the carrier gas (i.e., nitrogen 97.9 % pu­
rity). Then, the test gas (i.e., water vapor) was flushed. To remove any
influence caused by film thickness differences, transmission rate values
(g/m2⋅day) were converted into WVP values (g⋅mm/m2⋅day⋅KPa). WVP
was performed at least in duplicates for each film.

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Carbohydrate Polymers 294 (2022) 119780

Fig. 1. Degree of substitution (DStotal) of acetylated pea starch (APS) at increasing temperature and reaction times, together with the DStotal values obtained of
acetylated maize starch (AMS) at 135 ◦ C and increasing reaction times. DStotal was determined by back titration and reported in Table 1.

even at 130 ◦ C for 8 h (Imre & Vilaplana, 2020). The higher reactivity of
pea starch compared to maize starch could be explained by the different
effect of temperature on granule microstructure over the course of starch
derivatization, and, consequently, on granular reaction locale. Smooth
pea starch exhibits lower gelatinization temperature attributed to the Btype crystallinity located in the center of pea starch granules (Li et al.,
2019). During gelatinization, the melting of the crystalline structure is
initiated from the central hilum of starch granules, and the B-type

allomorph possesses a lower melting temperature than A-type counter­
part due to the loose packing of the former (Jane et al., 1999).
Furthermore, the absence of amylose lipid complexes of pea starch, as
opposed to normal maize starch, results in a significantly lower onset
temperature to paste (or swell) than maize starch (Ren et al., 2021). In
this regard, some granular swelling has been deemed necessary to ach­
ieve chemical derivatization (even at the surface) (Huber & BeMiller,
2001). It can be concluded that the extent of derivatization was not only
a function of the properties of the reaction medium, but of the
temperature-driven dissociation of double-helical crystallites, loss of
crystallinity and, eventually, promotion of granule swelling, which
presumably is more likely to occur at shorter times in C-type granules,
such as those from smooth pea, at those high temperatures to attain high
DS.

were used to obtain mechanistic understanding of the acetylation of pea
starch and accurately determine the DS value. Fig. 2b shows the over­
lapping of the FTIR spectra of the native (NPS) and acetylated (APS) pea
starch at 135 ◦ C and increasing reaction time (0.5 h to 4 h). NPS showed
the characteristic bands of polysaccharides, such as a broad band,
named a, centered around 3300 cm− 1 and due to the stretching vibra­
tion of O–H groups, a less intense band around 2932 cm− 1 due to the
stretching vibration of C–H groups, and a series of bands in the
fingerprint area of the spectrum at 1149, 1076, 994, and 923 cm− 1
corresponding to the coupling mode of the C–O and C–C stretching, CO-H bending mode, C–O bond stretching, and α-1,4 glycosidic linkage
vibration, respectively (Tupa et al., 2013; 2015). FTIR spectra of APS
contained new bands characteristic of the acetate groups. Specifically, a
– O stretching
band near 1740 cm− 1 (named b) corresponding to the C–
vibration of ester carbonyl groups; a band near 1367 cm− 1 (named c)

corresponding to the deformation vibration of the alkyl –CH3 band; and
an intense band at 1217 cm− 1 (named d) attributed to the C-O-C
stretching vibration of the ester moieties (Tupa et al., 2015). The
appearance of these spectral bands, together with a reduction of the
intensity of the band a due to -OH groups located along the anhy­
droglucose units, confirms the formation of starch acetates, even at the
shortest reaction time (0.5 h). The intensity of these bands increased
together with the reaction time. However, no increase in intensity was
observed after 2 h, apparently indicating that no further esterification
was achieved. It must be noted, however, that due to the ability of tar­
taric acid catalyst to form esters with starch, bands b and d (vibration of
ester groups) could also be dependent on the formation of tartaryl side
group, and not only alkanoyl groups from acetic anhydride. Similar to
back titration, FTIR was not able to distinguish carboxylic acid species
and related side groups. Therefore, starch esters were also studied by
NMR to specifically distinguish acetate from tartrate starch.
NPS and APS were firstly studied by Proton Nuclear Magnetic
Resonance (1H NMR). Special attention was paid to solubilize APS at
high DS since it presented poor solubility in DMSO and other organic
solvents (Fig. S1), in agreement with previous studies (Tupa et al.,
2020). This effect suggested that crosslinking involving a polycarboxylic

3.2. Degree and nature of substitution by spectroscopic techniques
The degree of substitution analyzed by chemical titration does not
distinguish between carboxylic acid species and related side groups, e.g.,
tartaryl side groups, which also results in overestimation of the DS of
alkanoyl groups (Nielsen et al., 2018; Volant et al., 2020). During the
acetylation reaction, not only the anhydride reagent can participate in
the substitution reaction, but also the dicarboxylic acid catalyst can
react to form starch tartrates, as illustrated in Fig. 2a. Back titration is a

non-specific method that does not differentiate starch acetate from
starch tartrate, resulting in an important overestimation of the deter­
mination DS of alkanoyl groups (Imre & Vilaplana, 2020). Thus, Fourier
Infrared Spectroscopy (FTIR) and Nuclear Magnetic Resonance (NMR)
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Fig. 2. a) Acetylation of pea starch mediated by the organocatalyst L-(+)-tartaric acid with the two possible products resulting from the reaction, starch acetate or
starch tartrate. It must be noted tartaric acid-starch crosslinking can occur with any of the available hydroxyl groups of the anhydroglucose units. b) FTIR spectra of
native (NPS) and acetylated (APS) pea starch at different reaction times. Band a (near 3470 cm− 1), b (near 1740 cm− 1), c (near 1367 cm− 1) and d (near 1217 cm− 1)
correspond to the -OH groups, ester carbonyls, alkyl -CH3 deformation, C-O-C stretching vibrations, respectively. c) Single Pulse-13C NMR spectra where signal 1
corresponds to the carbon of the alkyl group, signal 3 to the C2, 3, and 5 of starch, signal 2, 4 and 5 to the C6, C4 and C1, respectively, and signal 6 to the carbon of the
ester group.

acid, i.e., tartaric acid catalyst, could be occurring during the process, as
suggested by Tupa et al., 2020. To overcome this drawback, APS was
dissolved in DMSO‑d6 by heating at high temperature (over 100 ◦ C) for
at least 30 min until complete dissolution. Reference signals of the
protons from the anhydroglucose units were observed between 4.4 and
5.6 ppm due to the hydroxyl groups (-OH) of C2, C3, and C6 (Fig. S2). The
signal at 5.10 ppm is attributed to the anomeric proton of the (α-1,4)
linkage, whereas the small signal at 4.86 ppm corresponds to the
anomeric proton of the (α-1,6) linkage. The protons of the C2 to C6
anhydroglucose units resonate between 3.8 and 3.2 ppm (Kemas et al.,
2017). Esterified starch showed additional signals between 1.8 and 2.2
ppm due to the methyl protons of the acyl groups, demonstrating a

successful acetylation process (see Fig. S2) (Elomaa et al., 2004; Kemas
et al., 2017). The degree of acetylation calculated using Eq. (3),
increased over time reaching the maximum after 3 h of reaction
(Table 1). Values were significantly lower than those obtained by the
classical titration, confirming the overestimation of the titration method
due to interactions of the starch polymer with the catalyst. 1H NMR was
demonstrated to be a suitable and more accurate alternative than the
titration due to the specificity of the determination. However, the poor
solubility of the starch acetates, and the possible overlapping of some
signals with the residual solvent or water, challenges the accurate
determination of the degree of acetylation (Hadi et al., 2020). Hence,
solid state 13C-SP- and CP/MAS-NMR was also performed to determine
the degree of esterification independently achieved by acetate (DSacyl)

and by tartrate (DStar). NPS spectra (Fig. 2c) showed four main signals
due to the C1-C6 anhydroglucose units: signal 2 at 57.6 ppm corre­
sponding to C6, signal 3 at 68.3 ppm attributed to C2, C3 and C5, signal 4
at 77.8 ppm due to C4, and signal 5 at 96.8 ppm corresponding to C1
(Nielsen et al., 2018). In acetylated pea starch, two additional signals
appeared, namely signal 1 at 16.4 ppm and signal 6 at 166.6 ppm,
corresponding to the carbon of the methyl protons of the acyl group
(-CH3) and the ester group (C=O), respectively. The area of both signals
increased with the reaction time, concomitant with a significant
decrease of the intensity of signal 2 (C6) as well as signal 3 (partially due
to C2 and C3 of the anhydroglucose), demonstrating the acetylation in
these carbons (Nielsen et al., 2018). DSacyl and DStar were calculated
from the 13C SP-NMR spectra and compared to DS obtained by titration
and 1H NMR (Table 1). The total degree of acetylation (DStotal) increased
with the reaction time, reaching a maximum of ca. 3 after 3 h at 135 ◦ C
and agreeing with 1H NMR results. As expected, 1H NMR slightly

underestimated the total degree of substitution (due to the aforemen­
tioned challenges), although a significant correlation (p < 0.0001; R2 =
0.99) was observed between the liquid and solid-state NMR results. At
short reaction times (0.5 h), no reaction of the catalyst tartaric acid with
the starch was observed and thus, there were no significant differences
with the DS value calculated by titration (p > 0.05). However, after 1 h,
the catalyst competed with the acetic anhydride, resulting in up to 16 %
of the total esterification ruled by tartaric acid in 4 h (Table 1). NMR
data clearly confirmed that the DS by the chemical titration method is
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accurate and reliable when measuring at low DS values in organo­
catalytic reactions; nevertheless, the interaction of tartaric acid with the
hydroxyl groups of the anhydroglucose residues at longer reaction times
led to an overestimation of the DS values. This phenomenon agreed with
previous studies in which the contribution of tartaric acid in substitution
reactions with maize starch was demonstrated by HPLC-UV (Imre &
Vilaplana, 2020). Likewise, Nielsen et al. determined the degree of
butyrylation of maize and potato starches by 13C-SP- and CP/MAS-NMR
and reported overestimations between 38 and 91 % of the DS value by
the chemical titration (Nielsen et al., 2018). In our study, the derivati­
zation of pea starch was mainly ruled by acetylation (from 100 % at ≤1 h
reaction times to 84 % at reactions times up to 4 h) and not by tartrate
formation. The low reactivity of tartaric acid, and the efficient acetyla­
tion process of pea starch, were likely the result of several factors, such

as the type of acyl donor used (anhydride instead of acid), the low water
content of the starch sample due to the drying step prior to the reaction,
as well as the short carbon chain-length of the acyl donor reagent (Imre
& Vilaplana, 2020). It is worth noting that, due to the dihydroxy
dicarboxylic nature of tartaric acid, a possible formation of diacetyl
tartaric acid anhydride and further esterification of the acyl groups with
the starch cannot be ruled out (Tupa et al., 2020). Likewise, the dihy­
droxy dicarboxylic nature of tartaric acid could likely result in cross­
linked di-starch tartrate, which notably decreased its solubility in
organic solvents (Fig. S1).

acetylation of maize starch with classical NaOH (Xu et al., 2004) or
tartaric acid-catalyzed reactions at intermediate DS (Imre & Vilaplana,
2020), which has been suggested to be advantageous in the composites
of starch with synthetic polymers as this could improve interfacial
adhesion (Imre & Vilaplana, 2020).
Even in the absence of water, NPS showed a gradual loss of bire­
fringence at increasing reaction times, indicating a continuous breakdown of the crystalline structure during acylation. X-ray diffraction
patterns revealed the characteristic C-type polymorphism with peaks at
diffraction angles 2θ of 15.1◦ , 17.1◦ , 18◦ , and 23◦ (Fig. 3b). On one hand,
organocatalytic acetylation did not alter the number and position of the
diffraction peaks. On the other hand, it resulted in a progressive loss of
the intensity of all peaks, which followed a similar trend to that of
birefringence and evidenced a continuous breakdown of the crystalline
structure during acetylation. This occurrence has also been reported for
tartaric acid catalyzed maize starch using acetic anhydride as acyl
donor, which was attributed to the introduction of acyl groups during
the esterification reaction (Tupa et al., 2020) and to the potential plas­
ticizing effect of the side-product of the reaction, acetic acid (Imre &
Vilaplana, 2020). Interestingly, even high-DS anhydride-treated samples

(DSacyl ~ 3) retained their distinct granular structure (see APS-3 h in
Fig. 3a). This event could be the consequence of the formation of
crosslinked di-starch tartrate, as 13C-SP/MAS NMR data suggested
(Table 1), which would result in a cross-linked external shell that pre­
vents granular disruption, as suggested by Tupa et al., 2020.
The SEC-RI elution profile of NPS displayed two distinct peaks for
amylose and amylopectin molecules separated at ~14 mL elution vol­
ume, both represented by a single peak in the MALS detector that cor­
responded to a weight average molar mass for amylopectin molecules of
7.49 × 106 g/mol (Fig. 4a, Fig. S3). These results agreed with previous
studies using pea starch (Martinez et al., 2018). It is noteworthy that
with the sample concentration and injection volume used in this study
and amylose molecules being highly polydisperse, they do not exhibit
detectable laser-light scattering signals, as shown before (Roman et al.,
2019). Generally, tartaric acid catalyzed esterification caused a gradual
shift of the peaks towards higher elution volumes, confirming hydrolytic
degradation and a consequent molar mass decrease (Fig. 4a, Table 1).
Similar findings were already reported before for maize starch esterified
with acetic anhydride, which was explained by the effect of high tem­
peratures in acidic environment (Imre & Vilaplana, 2020). The reaction
also decreased the area under the peaks and resulted in a monomodal
size distribution, which could result from the coelution of amylose and
amylopectin fragments and the fact that low molar mass fractions might
be particularly hydrolyzed and washed out of the sample during the

3.3. Effect of tartaric acid catalyzed acetylation process on starch
granular morphology, birefringence, crystallinity, and molecular weight
(Mw)
Native pea starch granules exhibited oval, spherical, kidney and
irregular shape and a bimodal size distribution, although the large

population of granules (~40 μm of mean diameter) was significantly
more dominant (Fig. 3a). The observation of NPS granules under
polarized light exhibited the typical birefringence attributed to their
radial crystalline structure (Fig. 3a). Morphology and birefringence
were not altered during acetylation at reaction times ≤1 h (APS-0.5 h
and APS-1 h), i.e., starch acetates with DSacyl ≤ 1. Similar findings were
reported for organocatalytic esterification of maize starch at low DS
(Tupa et al., 2013). Nevertheless, acetylation at longer reaction times
resulted in a gradual increase of granular surface roughness, decrease of
birefringence, and the appearance of granular aggregation. At 3–4 h
reaction time (DSacyl ~ 3), fusion of granules, alongside a dramatic in­
crease in surface roughness and a complete loss birefringence were
detected. Increase of roughness upon acylation was also observed in the

Fig. 3. a) Visual appearance and morphology of native (NPS) and acetylated pea starch (APS) granules under the light (upper) and polarized light (lower) mi­
croscopy. Acetylation reaction temperature was 135 ◦ C and the granules were obtained after different reaction times that resulted in different DS. From left to right:
NPS, APS-0.5 h (DSacyl = 0.39), APS-1 h (DSacyl = 1.00), APS-2 h (DSacyl = 2.23), APS-3 h (DSacyl = 2.80), APS-4 h (DSacyl = 2.63). b) X-Ray diffraction patterns of
NPS and APS at different reaction times.
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Fig. 4. a) SEC elution profiles of native (NPS) and acetylated (APS) pea starch at increasing reaction times (0.5 h to 4 h) and constant reaction temperature of 135 ◦ C
obtained by HPSEC-MALS-RI. b) Thermal stability represented as weight loss (%) as a function of temperature (upper chart), as well as the derivative mass loss (lower
chart), determined by TGA. c) Thermograms showing the glass transition temperature (Tg) measured by DSC. APS-0.5 h (DSacyl = 0.39), APS-1 h (DSacyl = 1.00), APS2 h (DSacyl = 2.23), APS-3 h (DSacyl = 2.80), APS-4 h (DSacyl = 2.63).

purification. Interestingly, the rate of molar mass decrease with time

was significantly lower at longer reaction times, and even the weight
average molar mass increased from 3 to 4 h of reaction (Table 1). This
occurrence supports 13C-SP/MAS NMR data and the idea that tartaryl
esterification formed crosslinked di-starch tartrate due to the dihydroxy
dicarboxylic nature of tartaric acid.

was attributed to the release of acetic acid from anhydroglucose units
(Imre & Vilaplana, 2020; Thiebaud et al., 1997; Tupa et al., 2013).
Results showed that tartaric acid catalyzed acetylated pea starch of
DSacyl ≥ 2.6, whose lower temperature degradation step was completely
eliminated, presented an enhanced thermal stability despite their molar
mass decrease and loss of crystalline structure. The substitution of the
hydroxyl by alkanoyl groups in the modified starch seem to avoid interand intramolecular dehydration reactions ruling the decomposition of
pea starch.
We also investigated the effect of acetylation on the glass transition
temperature (Tg), which was revealed after a second heating run during
DSC analysis (Fig. 4c). Native pea starch (NPS) displayed a Tg of
110.8 ◦ C (Table 2). It is well known that the crystalline structure of
starch granules acts as physical cross-linking avoiding the mobility of
the polymer chains (Mizuno et al., 1998). In this regard, a decrease in Tg
upon acetylation was expected as a consequence of the gradual decrease
of crystallinity (Fig. 3b) and molar mass (Fig. 4a). However, Tg gradually
increased from 110.8 ◦ C (NPS) to 153.6 ◦ C (4 h reaction), which could be
explained by the increase in DS of tartaryl side groups (Table 1). Spe­
cifically, tartaryl esterification would reduce the molecular mobility,
either through strong hydrogen bonding of tartaric acid with other
starch chains hydroxyl groups (Imre & Vilaplana, 2020), or more likely
by the formation of covalent crosslinked di-starch tartrate, as suggested
by the SEC-MALS-RI molar mass distributions (Fig. 4a).
Overall, results demonstrated that tartaric acid catalyzed acetylation

of pea starch with acetic anhydride significantly increased its degrada­
tion temperature and Tg, a key factor to broadening its processing
temperatures.

3.4. Hydrophilicity and thermal stability of tartaric acid catalyzed pea
starch acetates
Starch hydrophilicity, one of its main shortcomings for many appli­
cations as biomaterial, was indirectly investigated from the first weight
loss shown in the TGA weight-loss and derivative mass loss curves
(Fig. 4b). Native pea starch depicted a first weight loss of ~10 % (be­
tween 45 and 140 ◦ C and a maximum peak at 85.5 ◦ C) due to the
evaporation of the remaining water bound to starch (Di Filippo et al.,
2016; Tupa et al., 2013). Interestingly, this loss was indirectly correlated
with DSacyl (p < 0.001; R2 = 0.983), demonstrating that starch hydro­
philicity was significantly reduced by acetylation.
The decomposition temperature of starch samples, and hence their
thermal stability and processability, increased with the degree of acet­
ylation. Native pea starch showed a single weight loss peak at a
maximum of 318.9 ◦ C. As acetylation time increased, a second weight
loss step at higher temperature appeared (369.3–374.5 ◦ C) with the
concomitant dissipation of the lower degradation step. This phenome­
non was also observed to occur during the tartaric acid catalyzed
esterification of maize starch (Elomaa et al., 2004; Imre & Vilaplana,
2020; Tupa et al., 2013). The first step corresponds to the condensation
of the remaining non-esterified -OH groups, whereas the second peak
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Carbohydrate Polymers 294 (2022) 119780

starches (Colussi et al., 2017; El Halal et al., 2017).

Table 2
Glass transition temperature (Tg) and thermal degradation (Td) of native (NPS)
and acetylated pea starch (APS) studied by DSC and TGA, respectively.

Tg (◦ C)*
Td1 (◦ C)
Area Td1
(mg/oC)
Td2 (◦ C)
Area Td2
(mg/oC)

NPS

APS-0.5
h

APS-1 h

APS-2 h

APS-3 h

APS-4 h

110.8

± 2.2c
318.9
± 0.6a
19.5 ±
0.1a
–

136.6
± 3.4b
313.7
± 0.9b
6.7 ±
0.2b
373.8
± 0.4ab
2.2 ±
0.1c

129.5
± 2.6b
289.2
± 2.4c
1.7 ±
0.1c
374.1
± 1.3a
9.4 ±
0.2b

130.3

± 2.9b
–

137.1
± 2.8b
–

153.6
± 5.6a
–

–

–

–

369.3
± 0.4c
21.3 ±
0.2a

374.5
± 0.7a
22.9 ±
0.3a

372.3
± 0.1ab
19.3 ±

3.9a

–

3.6. Thermal and mechanical properties and Water Vapor Permeability
The thermal decomposition of the biofilms studied by TGA revealed
four degradation steps attributed to the main components of the biofilms
(Fig. 6a). Biofilms showed a first peak corresponding to the remaining
bound water between 50 and 150 ◦ C, with a maximum peak near 80 ◦ C.
The weight loss % was ~6 % in NPS biofilms and a gradual loss decrease
occurred when using intermediate [BAPS-0.5 h (~4.7 %)] and highly
substituted APS [BAPS-1 h to -4 h (~4.1 to 4.5 %)]. This event evidenced
the enhanced moisture resistivity of APS biofilms compared to its NPS
counterpart. The first decomposition step of APS films was detected in
the range of 208 to 218 ◦ C (Td1), which was absent in NPS films
(Table 3). Since the decomposition of the glycerol plasticizer occurs
between 150 and 290 ◦ C (Sanyang et al., 2015), we believe this degra­
dation step corresponds to the degradation of glycerol that is poorly
interacting with starch in APS-based films. Specifically, the mass loss at
this temperature increased with the degree of substitution. A second
degradation step occurred in the range between 320 and 324 ◦ C (Td2),
which was attributed to the decomposition of native starch. Logically,
the area of this step decreased as DS increased, since technically there is
less non-esterified starch present in the films. No major differences were
detected in Td2 with the degree of acetylation. A third degradation peak
in the range between 364 and 368 ◦ C appeared in APS films (Td3), which
by all likelihood represents the decomposition of alkanoyl groups in the
modified starch (see Section 3.4). As previously reported, acetylation
limits intramolecular dehydration of polysaccharides by reducing the
hydroxyl groups (Aburto et al., 1999; Fundador et al., 2012), which, in

turn, delays the initiation of thermal decomposition.
The mechanical properties of APS-films were measured under ten­
sion, and the tensile strength (MPa), elongation at break (%), and
Young's Modulus (MPa) were determined from the stress-strain curves of
each film (Fig. 6b, Table 3). NPS films exhibited mechanical properties
similar to those reported in other studies for pea starch films (Cano et al.,
2014). The tensile strength and Young's Modulus significantly increased
when using APS at low degree of substitution (DSacyl = 0.39), compared
to native starch films. The reduction of intermolecular interactions
within the starch phase as a consequence of surface lamellar acetylation
in low DS starch could explain the alteration of the blend morphology
and decreased particle size (as observed in Fig. S4). We believe that this
occurrence could lead to a more homogeneous dispersion of the
dispersed phase. It is noteworthy that at short reaction time, APS
showed molar mass above 2.65 × 105 Da, which is significantly higher
than that reported for tartaric acid catalyzed maize starch using acetic
acid (Imre & Vilaplana, 2020). Therefore, the reduction in molar mass
could have played a minor negative role in APS-0.5 films. Nevertheless,
APS with higher DS showed a gradual decrease in the tensile strength
and Young's Modulus as DS increased (Table 3). The same mechanisms

*
Tg, glass transition temperature measured in a second heating cycle by DSC.
Td1 and Td2 represent the degradation peaks detected by TGA. Values are
expressed as average (n = 2) ± standard deviations. Values followed by different
letters within each parameter (row) indicate significant differences (p ≤ 0.05). -,
not detected.

3.5. Film-forming properties of tartaric acid catalyzed APS
Since tartaric acid catalyzed acetylation of pea starch significantly

enhanced its moisture resistance and thermal stability, we investigated
the role of an increasing degree of substitution on the film-forming
properties of APS. Firstly, freestanding biofilms with APS as the only
matrix polymer were made by solvent casting. Nonetheless, cohesive
films were not attained even using the acetylated pea starch with the
lowest DS (DSacyl = 0.39). Hence, NPS/APS blends (3:1 w/w) were
prepared instead, which allowed us to understand the effect of DS on
film properties, and the compatibility of APS with its native counterpart,
NPS. NPS films were clear, transparent, and presented a smooth surface
(Fig. 5a). When adding acetylated starch at DSacyl ≤ 0.4, films were still
transparent and smooth, although certain evidence for phase separation
was already visible. Biofilms became gradually and visibly rougher,
darker in color, less transparent and thicker as APS with higher DS was
used (Fig. 5b-f, Table 3). This could be explained by two factors. Firstly,
the substitution of -OH groups with hydrophobic monofunctional re­
agents would not only decrease inter- and intra-molecular interactions
within the polysaccharide phase, but it could also lead to less adhesion
at the interface between the composite components. This could result in
meaningful phase separation and deterioration in APS film properties
(Imre et al., 2019). In fact, sharp edges and large cavities were observed
around the starch granules in the films made with APS at high DS
(Fig. S4). Secondly, the decrease of molar mass during acetylation could
have lowered the availability of chains for matrix interaction. Likewise,
a darker color was expected due to the dark color that results from
acetylation (Fig. 3a). It is noteworthy that the low standard deviation in
film thickness evidenced the uniformity of the films. These values agree
with the thickness of other biofilms made with acetylated barley and rice

Fig. 5. Biofilms made of (a) native pea starch, and blends of native: acetylated (3:1, w:w) pea starch of (b) APS-0.5 h (DSacyl = 0.39), (c) APS-1 h (DSacyl = 1.00), (d)
APS-2 h (DSacyl = 2.23), (e) APS-3 h (DSacyl = 2.80), (f) APS-4 h (DSacyl = 2.63).

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Table 3
Mechanical, thermal and barrier properties of biofilms made with native pea starch (BNPS) or native/acetylated pea starch blends (BAPS) at a 3:1 ratio (w/w).
BNPS

BAPS-0.5 h

BAPS-1 h

BAPS-2 h

BAPS-3 h

BAPS-4 h

Mechanical properties
Thickness (mm)
Tensile Strength (MPa)
Elongation at break (%)
Elastic modulus (MPa)

0.09 ± 0.00c
17.8 ± 1.3b
4.3 ± 0.1ab

804.9 ± 14.4b

0.08 ± 0.00c
20.1 ± 1.7a
2.4 ± 0.7b
1003.5 ± 136.2a

0.10 ± 0.00b
13.9 ± 0.8c
3.1 ± 0.3b
689.3 ± 83.0c

0.11 ± 0.00a
7.1 ± 0.4d
5.4 ± 0.7a
332.5 ± 43.6d

0.11 ± 0.02ab
6.4 ± 0.9d
5.6 ± 0.5a
263.6 ± 25.7d

0.12 ± 0.01a
5.8 ± 0.3d
3.8 ± 0.5b
302.7 ± 10.8d

Thermal stability (TGA)
Td1 (◦ C)
Area Td1 (mg/ oC)

Td2 (◦ C)
Area Td2 (mg/ oC)
Td3 (◦ C)
Area Td3 (mg/ oC)

–
–
320.6 ± 2.9b
17.6 ± 2.2a
–
–

208.8 ± 2.6b
1.3 ± 0.1d
323.1 ± 0.1ab
11.6 ± 0.3b
368.3 ± 0.8a
0.7 ± 0.1c

218.8 ± 0.5a
1.5 ± 0.0cd
323.5 ± 0.5ab
11.0 ± 0.2b
367.1 ± 0.6ab
1.7 ± 0.0a

207.1 ± 6.7b
1.6 ± 0.0bc
324.1 ± 0.4a
10.1 ± 0.7b

367.1 ± 0.4ab
1.7 ± 0.1a

214.5 ± 0.0ab
1.8 ± 0.1ab
324.2 ± 0.5a
9.6 ± 0.0b
365.7 ± 0.7b
1.4 ± 0.1b

214.3 ± 1.9ab
2.0 ± 0.0a
322.2 ± 0.7ab
10.7 ± 0.4b
364.0 ± 0.5c
0.4 ± 0.1d

Water Vapor Permeability (WVP)
WVP (g⋅mm)/(m2⋅day⋅KPa)

1.03 ± 0.07ab

0.74 ± 0.09c

0.92 ± 0.00b

1.11 ± 0.02a

0.93 ± 0.01b

1.02 ± 0.08ab

Values followed by different letters within each parameter (row) indicate significant differences (p ≤ 0.05). Td1, Td2, and Td3 represent the degradation temperature
peaks detected by TGA. -, not detected. BAPS-0.5 h (DSacyl = 0.39), BAPS-1 h (DSacyl = 1.00), BAPS-2 h (DSacyl = 2.23), BAPS-3 h (DSacyl = 2.80), BAPS-4 h (DSacyl =
2.63).

Fig. 6. a) Thermal stability of native (BNPS) and acetylated pea starch (BAPS) biofilms, made with acetylated pea starch at different DS, represented as weight loss
(%) as a function of temperature (left), as well as the derivative mass loss (right), determined by TGA. b) Strain stress curve of BNPS and BAPS biofilms. APS-0.5 h
(DSacyl = 0.39), APS-1 h (DSacyl = 1.00), APS-2 h (DSacyl = 2.23), APS-3 h (DSacyl = 2.80), APS-4 h (DSacyl = 2.63).

governing film roughness and transparency (see Section 3.5) could also
explain mechanical properties. Schmidt et al. also reported improved
tensile strength at low DS values and poorer mechanical resistance at
high DS investigating NaOH catalyzed acetylated cassava starch biofilms
(Schmidt et al., 2019). The elongation at break (EB%) using APS DSacyl

≤ 1 was similar or slightly worse than that of NPS films (Fig. 6b,
Table 3), in agreement with other low acetylated starch films (L´
opez
et al., 2011; Schmidt et al., 2019). Nonetheless, EB% improved when
APS DSacyl > 2 was used in comparison with the control NPS film. The
retained crystallinity of low acetylated starch probably makes these
10


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Carbohydrate Polymers 294 (2022) 119780

films more brittle and less flexible than highly acetylated starch films in

which the crystallinity was lost. Interestingly, a gradual decrease in the
EB% was observed from 2 h to 4 h, which was likely the result of the
reduction of molecular mobility and increase in rigidity due to tartaryl
esterification and/or crosslinked di-starch tartrates formation.
The Water Vapor Permeability (WVP) of the native and APS films
was measured to assess their potential as barrier materials (Table 3).
NPS films possessed a WVP of 1.03 ± 0.07 g⋅mm /m2⋅day⋅KPa, which is
in accordance with other values reported for starch-based films (El Halal
et al., 2017). The incorporation of APS-0.5 significantly reduced the
WVP of the films due to the reduced hydrophilicity of starch acetates
(see Section 3.4). Nonetheless, highly acetylated pea starch films (DSacyl
≥ 1) did not show significant differences (p > 0.05) with the control,
which, by all likelihood, is the consequence of phase separation resulting
in a heterogenous and microporous structure from which water vapor
can permeate (Fig. S4).

Methodology. Mingwei Geng: Investigation, Methodology. Mario M.
Martinez: Conceptualization, Methodology, Supervision, Project
administration, Funding acquisition, Formal analysis, Visualization,
Writing – review & editing.
Declaration of competing interest
The authors declare that they have no known competing financial
interests or personal relationships that could have appeared to influence
the work reported in this paper.
Acknowledgments
This research was funded by Aarhus University Research Foundation
(Aarhus Universitets Forskningsfond, AUFF), project number AUFF-F2020-7-5. The authors would like to thank Laura Roman for her assis­
tance in the analysis of starch molecular weight, as well as Thomas
Vosegaard and Dennis Wilkens Juhl for their assistance in the solid-state
NMR analysis. The authors also thank Cosucra which generously sup­

plied the raw pea starch.

4. Conclusions
In this study, we systematically reported the organocatalytic acyla­
tion of smooth pea starch using tartaric acid as green catalyst and acetic
anhydride as acyl donor. Pea starch was less recalcitrant towards
organocatalytic esterification than maize starch, as indicated by the
faster reaction kinetics and higher degree of substitution observed in the
former. The degree of substitution with alkanoyl (DSacyl) groups was
validated using FTIR, 1H NMR and solid-state 13C NMR. However, only
NMR displayed good specificity and 13C-SP-NMR enabled the quantifi­
cation of the degree of substitution by tartaryl groups (DStar). Pea starch
with DSacyl from 0.03 to 2.8 was successfully developed, with granular
and crystalline structure mostly retained until DSacyl ≤ 1.0. Longer re­
action time resulted in starch granule surface roughness, loss of bire­
fringence, and hydrolytic degradation. Reaction time and temperature
played a key role to attain high DSacyl, although high reaction times at
high temperature resulted in up to 16.2 % of the overall esterification
with tartaric acid (DStar up to 0.5). 13C-SP-NMR, SEC-MALS-RI and the
significant decreased of solubility in organic solvents, suggested that
tartaryl groups participated in the formation of crosslinked di-starch
tartrate, which seems logical considering the dihydroxy dicarboxylic
nature of tartaric acid. Acetylation significantly increased the hydro­
phobicity, thermal resistivity and processability (broader processing
temperature) of pea starch. Moreover, the use of organocatalyticallyacetylated pea starch with low DS (DSacyl ≤ 0.4) generated starchbased biofilms with higher tensile properties and lower water vapor
permeability, whereas high DS (DSacyl ~ 2) increased the elongation at
break of the films. Low performance on acetylated pea starch with high
DSacyl was attributed to the incompatibility between highly acetylated
and native pea starches.
The efficient organocatalytic esterification process shown in this

study resulted in pea starch acetate polymers with controlled degree of
substitution that could serve to replace base-catalyzed acetylated starch
esters typically used for food and biomaterial applications. This com­
plementary mode of catalysis has enormous potential for savings in cost,
time, and energy, resemble an easier experimental procedure, and
reduce chemical waste. Results from this study, together with the in­
clinations for clean labels, healthy eating, sustainability, and conve­
nience, could also fuel the growth of the global market for pea starch,
with still unpaired abundance and demand. Nonetheless, further studies
would still be needed to investigate the compatibilization of tartaric acid
catalyzed pea starch with other polysaccharides under scalable melt
mixing (e.g., extrusion) to enhance interfacial adhesion and subsequent
biomaterial properties.

Appendix A. Supplementary data
Supplementary data to this article can be found online at https://doi.
org/10.1016/j.carbpol.2022.119780.
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