Enzyme entrapment in amphiphilic myristyl-phenylalanine hydrogels

Article abstract of DOI:10.3390/molecules24162884

Supramolecular amino acid and peptide hydrogels are functional materials with a wide range of applications, however, their ability to serve as matrices for enzyme entrapment have been rarely explored. Two amino acid conjugates were synthesized and explore

Full text of DOI:10.3390/molecules24162884

AErnticlze yme Entrapment in Amphiphilic Myristyl-
Phenylalanine Hydrogels
1,2
2,3
2
1,2,3,
Natashya Falcone , Tsuimy Shao , Roomina Rashid and Heinz-Bernhard Kraatz
*
1
Department of Chemical Engineering and Applied Chemistry, University of Toronto, 200 College Street,
M5S 3E5, Toronto, Canada
Department of Physical and Environmental Sciences, University of Toronto Scarborough, 1065 Military
Trail, M1C 1A4, Scarborough, Canada
2
3
Department of Chemistry, University of Toronto, 80 St. George Street, M5S 3H6, Toronto, Canada
* Correspondence: Bernie.kraatz@utoronto.ca; Tel.: 416-287-7197
Academic Editors: Francesca D’Anna, Salvatore Marullo and Carla Rizzo
Received: 3 July 2019; Accepted: 5 August 2019; Published: 8 August 2019
Abstract: Supramolecular amino acid and peptide hydrogels are functional materials with a wide
range of applications, however, their ability to serve as matrices for enzyme entrapment have been
rarely explored. Two amino acid conjugates were synthesized and explored for hydrogel formation.
These hydrogels were characterized in terms of strength and morphology, and their ability to entrap
enzymes while keeping them active and reusable was explored. It was found that the hydrogels
were able to successfully entrap two common and significant enzymes—horseradish peroxidase
and α-amylase—thus keeping them active and stable, along with inducing recycling capabilities,
which has potential to further advance the industrial biotransformation field.
Keywords: amino acids; protein; hydrogels; enzyme entrapment
1. Introduction
Amino acid and peptide-based supramolecular gels are functional materials that have been the
subject of research due to their versatility and their wide range of applications in many fields [1].
Interactions generating the supramolecular network present in physical gels are the result of non-
covalent interactions between gelator molecules. Non-covalent interactions present in amino acid and
peptide gel materials include hydrogen bonding, van der Waals interactions, π−π stacking, and
hydrophobic interactions. Physical gels are also affected by the external environment and are able to
undergo sol–gel phase transitions in response to temperature [2], pH [3], and solvent, which allows
for tunable properties of these materials. Although supramolecular amino acid and peptide
hydrogels formed by self-assembly of amphiphilic molecules have served as scaffolds for tissue
engineering and wound healing [4,5], mediums for environmental remediation[6,7], biomaterials for
drug delivery, and as antimicrobial agents [8–10], their applications as matrices for enzymatic
reactions to occur in have been rarely explored.
Enzymes are a class of proteins that exhibit high efficiency and specificity for catalyzing many
reactions in our bodies and in biochemical and chemical reactions. In industrial biotransformations,
using enzymes as biocatalysts are attractive as they possess high selectivity, run under milder and
greener reaction conditions, and eliminate the need of expensive and toxic metal catalysts [11].
Enzymes are attractive “green” alternatives to chemical catalysts within the industrial sector, but
their robustness to environmental conditions needs optimizing. The issue with adopting enzymes as
biocatalysts on the industrial scale also includes their low stability and poor recycling ability.
However, it has been reported that immobilized enzymes have a greater stability compared to free
enzymes. There are three main immobilization methods: Adsorption [12], entrapment [13], and
Molecules 2019, 24, 2884; doi:10.3390/molecules24162884
www.mdpi.com/journal/molecules

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covalent bonding (Scheme 1) [14]. While all have pros and cons, entrapment, meaning the enzyme
remains within the gel matrix, has been more readily explored due to adsorption being weak and
covalent bonding resulting in binding of the active sites of enzymes. Most reports of enzyme
immobilization using the entrapment method commonly use silica or polymer matrices [15–22]. Chen
et al. encapsulated triacylglycerol lipase within biomimetic silica through the catalysis of
polyallylamine and found encapsulation not only stabilized the lipase but also enhanced the activity
[16]. A N,N’-methylenebisacrylamide (MBA)-cross-linked N-isopropylacrylamide (NIPAAm)
hydrogel was also used to form a hemin-based mimetic enzyme with a higher activity compared to
hemin alone [23]. In addition, numerous applications of alginate gel matrices have been reported in
enzyme entrapment studies [24,25].
Scheme 1. The three enzyme immobilization methods; adsorption, entrapment, and covalent bonding
(from left to right).
Entrapment in an amino acid or peptide supramolecular hydrogels are rarely reported but has
the potential to be one of the leading matrices for keeping the enzymes stable and active in an
aqueous, physiological material [26]. It also has the potential to be an improved matrix compared to
polymeric or alginate entrapment matrices due to the common characteristics peptide hydrogels have
with enzymes. These include amino acids as the main building block, supramolecular interactions as
the driving force for self-assembly, amphiphilicity of the structures, and a supramolecular structure
as the result of the formation of a network of nanofibers [26]. Park et al. used a diphenylalanine
hydrogel to immobilize a glucose oxidase enzyme for employment of a biosensing platform [27]. Xu
et al. reported a molecular peptide hydrogel-immobilized enzyme exhibiting supractivity and high
stability [20]. In addition, this group showed a supramolecular-hydrogel-encapsulated hemin as an
artificial enzyme to mimic peroxidase and exhibited self-immobilization of a phosphatase enzyme in
a gel for high activity and stability [26,28–31]. Miller et al. also reported enzyme entrapment within
a hydrogel matrix and described how molecular hydrogels are ideal natural materials for the support
of enzymes as they reversibly self-assemble under aqueous conditions at physiological temperatures
[32]. However, to avoid enzyme leaching they had to covalently attach an enzyme to a β-sheet self-
assembling peptide. Since then peptide hydrogels have been rarely explored as an entrapment matrix
compared to silica or polymer matrices. To the best of our knowledge there are no recent (<five years)
references of amino acid and peptide hydrogels for enzyme entrapment, but many reported for silica
alginate, or polymeric matrices. For these reasons the uses of amino acid and peptide hydrogels
specifically, as a matrix is significant as they possess many advantages including biocompatibility
and biodegradability, in addition the amphiphilic nature of the peptide nanofibers is also reported to
facilitate the transportation of the substrate and product through the hydrogel before and after
catalytic transformations [32].
In this article, we expand the uses of amphiphilic-amino acid hydrogels for one of the enzyme
immobilization methods as an entrapment matrix for potential improvement to their stability,
protection from degradation, and increase in the ability of enzyme recycling over longer periods of
time. Two amphiphilic amino acid conjugates composed of an aliphatic chain from myristic acid and
L- and D-Phenylalanine (Phe) have been synthesized and explored for hydrogel formation.
Previously, our group reported only the L enantiomer of the compound in a 0.1 M phosphate buffer
solution at pH 7.25–8.5 and looked at metal binding studies [33]. Here, both enantiomeric compounds
formed a hydrogel in a 50 mM monobasic phosphate buffer solution at physiological pH 7 and their
full characterization including their strengths through rheology experiments and morphology

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through transmission electron microscopy (TEM) studies was compared. Interestingly, it is also
shown in the CD spectrum that the Phe compound was found to form an ordered structure exhibiting
a distinct ultra-narrow peak around 220–230 nm, which attributes to the formation of Phe-based
chiral structures that correspond to structural aggregates.
Scheme 2. Scheme summarizing the overall research work. (a) Synthesis of amino acid conjugates
using Myristic acid (MA) and Phenylalanine (Phe); (b) formation of hydrogel through supramolecular
assembly of amino acid conjugates; (c) enzyme entrapped in supramolecular hydrogel, substrate (blue
circle) diffusing in and product (green triangle) diffusing out.
Both hydrogels were tested for entrapment capabilities with two common and significant
enzymes: Horseradish peroxidase (HRP) and α-amylase. Product quantification and recycling
experiments were performed, and it was found that the enzymes immobilized in the gels were stable
and active for one month and were able to catalyze multiple reactions over a one-week time period
with minimal leaching. Scheme 2 summarizes the work in this report. Here, we present a qualitative
evaluation rather than a quantitative kinetic evaluation at present to stay within the scope. The
synthesized hydrogels did not deactivate the enzyme and were found to support both enzymes to
perform catalytic reactions through the matrix.
2. Results
2.1. Synthesis of Myristic Acid-L/D-Phenylalanine (MA-L/D-Phe-OH)
MA- -Phe-OH was synthesized using standard carbodiimide coupling procedures using
L/D
myristic acid and the amino acid phenylalanine, in the presence of HOBt [34]. Further synthetic
details can be found in the experimental section. Chemical structures of the amino acid conjugates
are shown in Figure 1 and their full characterization can be found in the SI.

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Figure 1. (a) Aliphatic-amino acid hydrogel shown through an inverted vial test and the molecular
structure of MA- -Phe-OH; and (b) aliphatic-amino acid hydrogel shown through an inverted vial
test and the molecular structure of MA- -Phe-OH.
L
D
The self-assembling behavior of the conjugates was explored in a 50 mM monobasic phosphate
buffer solution at pH 7. When a hot solution of the compound cooled to room temperature, a
viscoelastic gel was obtained. Surprisingly, this compound had a very low MGC, meaning only a
small amount of compound is required to form the hydrogel material, thus making it cost- and
material-efficient. At this MGC however, the melting temperature of the gel is low at around 37 °C,
at which the gel turns into a liquid state. As we increase the concentration of gelator, the melting
temperature of the gel increases as more heat is required to break the increased amount of interactions
between more gelator molecules, therefore at our working concentration of 1% the Tgel was 51 °C. The
tan δ values will be discussed in the following section. The gelation information for the compounds
including the minimum gelation concentration (MGC), melting temperature of the gel (Tgel) and the
tan δ values at 1% is reported in Table 1.
Table 1. MGC, Tgel, and Tan δ (all at 1%) values of hydrogels in 50 mM monobasic phosphate buffer
at pH 7.
°
T
gel ( C)
Tan δ (Gel-)
Tan δ (Gel +
HRP)
0.43 ± 0.07
0.43 ± 0.02
Tan δ (Gel +
Amylase)
0.42 ± 0.12
0.42 ± 0.07
Compound
MA- -Phe-OH
MGC (w/v)
(MGC/1%)
L
0.25%
0.25%
37/51
37/51
0.41 ± 0.03
0.41 ± 0.11
MA- -Phe-OH
D
2.2. Rheology Studies
Frequency sweep rheology experiments were performed to study the mechanical strength of the
gels (Figure 2). In oscillatory shear measurements, the shear storage modulus, G’, loss modulus, G”,
and loss factor, tan δ, are critical hydrogel properties monitored against time, frequency, and strain.
The complex modulus of the tested material is G*(w) = G’ + iG” as the real (elastic or in phase) and
imaginary (viscous or loss or out of phase) components of G* [35]. For a typical viscoelastic gel, the
storage modulus, G’, remains higher than the loss modulus, G” [35,36]. G’ measures the deformation
energy stored during the shear process of a test material (i.e., stiffness of the material) and G” is

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representative of the energy dissipated during shear (i.e., the flow or liquid-like response of the
material) [35]. A piece of the 1% hydrogel material was placed under the rheometer geometry and
frequency sweep experiments were performed at 0.1% strain from a frequency range of 0.1–100 rad/s.
From the data shown in Figure 2 a and c, both hydrogels proved to be a viscoelastic gel material as
the G’ was greater than G”. The elastic modulus, E’, was estimated as E’ = 2G’(1 + ν), where G’ is the
storage modulus measured by the rheometer, ν is Poisson’s ratio assumed to be 0.5 [37], for M-F(L)
and (D) E’ was roughly found to be ~725 and 715 Pa. The tan δ value (G”/G’) is a representative of gel
elasticity that measures the energy loss owing to viscous flow compared to the energy stored. When
this value is greater than 1 the sample behaves as a viscous liquid; conversely, when the value is less
than 1 the sample behaves more as an elastic solid [35,36]. The tan δ values for both compounds are
reported in Table 1 and it was found that the L and D enantiomer compounds had the same tan δ
value of 0.41. The compounds also demonstrated shear-thinning properties.
Figure 2. (a) Frequency sweep experiments on MA-
L-Phe-OH. G’ (storage modulus) remains higher
than G” (loss modulus). (b) Step strain experiments of MA-L-Phe-OH. Cycle 1 under 0.1% straight
and G’ remains higher than G”, cycle 2 under 100% strain gel is reverted to the sol phase where G” is
higher than G’, cycle 3 lifts the strain and is back to 0.1% and is back in gel phase as G’ is greater than
G”. This is repeated for two more cycles to show self-healing properties of the gel. (c) Frequency
sweep experiments on MA-D-Phe-OH. G’ (storage modulus) remains higher than G” (loss modulus).
(d) Step strain experiments of MA-D-Phe-OH. Cycle 1 under 0.1% strain and G’ remains higher than
G”, cycle 2 under 100% strain gel is reverted to the sol phase where G” is higher than G’, cycle 3 lifts
the strain and is back to 0.1% and is back in gel phase as G’ is greater than G”. This is repeated for
two more cycles to show self-healing properties of the gel.
Step-strain experiments using the rheometer demonstrating thixotropic properties were also
performed in which strain was applied to the gel causing the gel to revert to the solution phase.
Thixotropic properties of the gel demonstrate that under normal conditions, the property is a gel and
under stressed conditions such as heat, force, or sonication, it becomes less viscous, however, when
the stress is removed the gel-state reforms. Self-healing gels that can restore their functionalities and
structures after damage have been developed as ‘smart’ soft materials [38]. Rheological studies show
the behavior of the gel under mechanical stress and the variation of the storage and loss modulus
have been studied under the application and release of shear stress. Stress causes the gel to break,
while release of the mechanical stress allows reformation of the hydrogel [39]. In the first cycle in
which 0.1% strain was applied, G’ is greater than G” confirming gel formation, whereas in cycle 2,
100% strain is applied and G” is found to be greater than G’, thus reverting it to sol state. Strain was
removed and set back to 0.1% in the third and fifth cycles to see if the gel reformed, interestingly,
both compounds exhibited thixotropic properties shown in Figure 2 b and d.

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2.3. Circular Dichroism
As amino acids represent one of the simplest classes of small biological molecules that are also
chiral, they were the first molecules to be characterized by circular dichroism (CD) [40]. To further
investigate the supramolecular structure and the presence of both enantiomers, CD spectroscopy was
performed. CD spectroscopy has been reported to be a valuable technique that can be used to
determine the molecular chiral arrangements in a self-assembled state [41,42]. Bands below 300 nm
arise from chiral amino acids, while L amino acids exhibit a broad positive CD peak in the 200–207
nm region. Typical CD peaks for amino acids result from the carbonyl bond at 190–200 nm
representing the π-π* transition and at 210–230 nm representing the n-π* [40]. One of the most well-
known properties of aromatic moieties, such as the phenyl ring in the amino acid phenylalanine
(Phe), is their ability to aggregate/stack through π−π interactions. Although single amino acids are
not known to form secondary structures, the amino acid phenylalanine has been highlighted in
literature as a special case as it resembles the self-assembling process of the diphenylalanine
compound [40]. Phe was found to form an ordered structure exhibiting an ultra-narrow peak around
220–230 nm, which attributes to the formation of Phe-based chiral structures that correspond to
structural aggregates [40]. As shown in Figure 3, the presence of chiral amino acids and the interesting
characteristic of the self-assembling Phe is highlighted and observed as a major peak at around 225
nm for the L-enantiomer is shown, while the D-amino acid induced the opposite helicity, revealing
that amino acid chirality is opposite for each enantiomeric pair. Smaller peaks are observed around
200 nm indicating the carbonyl bond, which, as mentioned, is typical of amino acids.
Figure 3. Circular dichroism (CD) spectra of MA- -Phe-OH in blue and MA-D-Phe-OH in green
L
showing both enantiomers being mirror images of each other in 1% gel state. The spectra also
demonstrate a narrow peak and the self-assembling characteristics of phenylalanine as reported in
the literature.
2.4. Morphological Investigation
To investigate the morphological features of the self-assembled amino acid conjugates,
transmission electron microscopy (TEM) studies were performed. The Formarv carbon coated copper
TEM grid was dipped into the hydrogel, followed by drying at room temperature and under vacuum.
The grid was then directly put on the TEM instrument mentioned in the experimental section. The
morphology of a nanofiber matrix is reported for both L and D (Figure 4), where we observed long
fibers, which showed a three-dimensional interweaving architecture. Fibril structures have been
reported as the main morphology of strong peptide based-gel materials [43–46]. The fibers had an
average thickness of 0.056 ± 0.011 μm for MA-L-Phe-OH and 0.058 ± 0.021 μm for MA-D-Phe-OH,
which is similar to reported amino acid and peptide hydrogels. A histogram of the fiber thickness for
both compounds is found in Figure S9. Additional TEM images of the hydrogels with entrapped
enzyme can be seen in Figure S10. A three-dimensional fibrous network is also observed with

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additional black particles within the fibers, one may speculate this small difference may be due to the
entrapped enzyme.
Figure 4. TEM images of MA-
L
-Phe-OH and MA-D-Phe-OH. (a–c) Three-dimensional fibrous network
of MA- -Phe-OH, and (d–f) three-dimensional fibrous network of MA-
L
D-Phe-OH. The scale bar is 2
μm for all images except b), which is 5 μm.
2.5. Enzyme Entrapment
To investigate the ability of our amino-acid hydrogels to act as a matrix for enzyme entrapment,
50 μL of a 1 mg/20 mL enzyme solution of both horseradish peroxidase (HRP) and α-amylase were
added to the hydrogels. Scheme 3 shows the reactions both enzymes catalyze. Specifically, this was
done by heating the compound in 50 mM monobasic phosphate buffer pH 7 until a clear solution was
obtained. Once the temperature of the vial cooled, 50 μL of the enzyme solution was added into the
gel solution. As the solution further cools to room temperature on the bench top, the compound forms
a hydrogel with the enzyme entrapped inside. With the addition of the enzymes, the resulting
hydrogels were determined to have similar viscoelastic properties as the tan δ values remained the
same for both L and D gels with both enzymes (Table 1). The rheology data for the gels with
entrapped enzymes can be seen in Figure S11.
Scheme 3. Enzyme reaction schemes of horseradish peroxidase (HRP) and amylase enzymes. HRP
converts pyrogallol (substrate) to purpurogallin (product) and amylase converts starch (substrate) to
maltose (product).
The substrate solution containing 2.6 mL of 50 mM monobasic phosphate buffer pH 7, 0.16 mL
of a 30% H
2
O2
solution, and 0.32 mL of a 50 mg/mL pyrogallol solution was added on top of the
enzyme-entrapped hydrogel. Within seconds, a color change was observed in the substrate solution

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indicating the formation of product, purpurogallin, which could be quantified using ultraviolet (UV)
-1
-1
spectroscopy at 420 nm and its extinction coefficient of 2640 M cm in 50 mM phosphate buffer at
pH 7. In Figure 5a, the color change of the substrate solution after 30 seconds was observed, indicating
that the enzyme was very active in our hydrogel. The color change was first noticed at the surface of
the gel and as time increases the color change throughout the whole gel was observed. This is due to
the initial contact of the substrate and enzyme on the hydrogel surface, however as the substrate
diffused in the gel and reacted with more enzyme, more product was formed causing the full gel to
turn the brown color as seen in Figure 5 a after one hour. The formation of product was then
quantified over a period of time as shown in Figure 5b–c. It is observed that as time increases, giving
the substrate time to diffuse into the gel and react with the entrapped enzyme, the formation of
product also increased. This follows a similar trend to the enzyme in solution, where as time
increased, product formation also increased. However, due to the instant and direct contact of the
enzyme with the substrate in solution, product formation plateaus much quicker. The activity of the
enzyme in solution can be found in Figure S12.

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Figure 5. a) Image of the amino acid hydrogels with HRP entrapped prior to substrate solution
addition on top and after three minutes. b) Quantification data of product formation using MA-L-
Phe-OH entrapped hydrogel over time using absorbance at 420 nm and the known extinction
coefficient of 2640 cm-1 M-1. c) Quantification data of product formation using MA-D-Phe-OH
entrapped hydrogel over time using absorbance at 420 nm and the known extinction coefficient of
2640 cm-1 M-1. d) Image of the amylase-entrapped amino acid with the substrate solution shown prior
to addition on top of hydrogel and after 10 minutes on top of the hydrogel and 15 minutes in boiling
water. When in contact of a reducing sugar, the color reagent containing dinitrosalicylic acid goes
from a yellow-orange color to a darker orange-red color. e) Quantification data of product formation
using MA-L-Phe-OH entrapped hydrogel over time using absorbance at 540 nm and the calculated
extinction coefficient of 1340.6cm-1 M-1. f) Quantification data of product formation using MA-D-Phe-
OH entrapped hydrogel overtime using absorbance at 540 nm and the calculated extinction coefficient
of 1340.6 cm-1 M-1.
The substrate solution for the amylase enzyme, consisting of 1 mL of a 100 mg/10 mL solution
of starch, 1 mL of 50 mM monobasic phosphate buffer pH 7, and 1 mL color reagent (dinitrosalicylic
acid), was placed on top of the hydrogel containing the amylase enzyme. Since maltose cannot be
directly quantified, a color reagent solution consisting of dinitrosalicylic acid was used as it is a well-
known method for quantification of reducing sugars [47,48]. In the presence of reducing sugars, such
as the product of this reaction (maltose), the yellow solution turns red. With a standard curve of
maltose and color reagent (Figure S13), the amount of product produced from the enzymatic reaction
was quantified. Figure 5d shows the substrate solution with a color reagent before being placed on
top of the enzyme-entrapped hydrogel, left for 10 minutes on top of the hydrogel, and then followed
by 15 minutes in boiling water after which the substrate solution turned red. This proves again that
our enzyme was active in the hydrogel as product formation is observed. The product was quantified
using the extinction coefficient found from the standard curve shown in Figure S13 and in Figure 5e–
f, an increase in product formation can also be seen over time.
2.6. Enzyme Leaching
When testing for enzyme entrapment it is important to look at enzyme leaching to ensure all of
the enzyme is not leaching out and catalyzing the reaction in solution on top of the gel, but that the
substrate is diffusing through the porous gel and reacting with the entrapped enzyme. In almost all
cases of entrapment, enzymes may leach out of the gel. It has been reported however, due to the non-
covalent forces between the amino-acid conjugates involved in supramolecular gelation, and the
same forces involved in an enzyme’s supramolecular structure, that these forces help stabilize most
of the enzymes in the gel [23]. To prove that the activity reported from the enzymes in the gel was
not solely due to enzymes leaching out and catalyzing the reaction in the solution on top of the gel,
leaching experiments were performed. These experiments involved placing only a phosphate buffer
on top of the gels with entrapped enzymes and taking aliquots of the buffer at different time points.
The aliquots were then added to the substrate solutions (same amounts used when testing the activity
of enzymes in the gel) and activity was measured. Further experimental detail can be found in the
experimental section. Figure S14 shows a comparison of activity of the product formation when
catalyzed on the gel with entrapped enzyme to the activity of product formation using the leached-
out enzyme in the buffer. It is clearly shown that although minimal enzymes are leaching and
catalyzing the reaction slowly, it does not compare to the amount of product formed from the
catalytic reaction through the gel. In addition, Figure S15 shows an image of two sets of L and D
hydrogels, one contains the enzyme inside the gel while the other does not, the enzyme is added later
to the substrate solution on top. Here, we can visually see that the substrate is being catalyzed in the
gel as a brown color developed for the hydrogels with enzyme entrapped, while no color change is
observed for the gel with no enzyme entrapped but rather just on top with the substrate solution. For
both enzymes in both L and D hydrogels, one can conclude that the activity is not solely coming from
enzyme leaching out and catalyzing the reaction in solution, but in fact the catalytic reaction is
occurring through the gel matrix with the majority of enzyme entrapped in the fibers.

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2.7. Enzyme Recycling and Stability
Reported advantages of enzyme entrapment in amino acid and peptide hydrogels are to increase
its stability over longer periods and to be able to reuse and recycle the same enzyme for many
catalytic reactions [26]. A recycling experiment was performed where the enzyme-entrapped
hydrogels catalyzed a fresh substrate solution every day for one week. Each day the substrate
solution was removed, and a fresh solution was placed on top of the hydrogel to see if the enzymes
were still active and could catalyze multiple reactions over the one-week period of time. Figure 6 a–
d shows the activity after 10 minutes in the enzyme-entrapped hydrogels every day over a one-week
period. Over time less product was produced, which was expected; however, the enzyme was able
to remain active and be re-used multiple times in the hydrogel. Here, the same entrapped enzyme
was able to catalyze multiple catalytic reactions without the usual requirement of a new catalyst each
day.
In addition to being able to recycle and reuse the enzyme over time, a preliminary long-term
stability test of the enzyme was performed. Here, the enzymes were placed in both L and D hydrogels
and were left alone for one month. After this time, the activity of the enzyme was tested by placing
fresh substrate solution on top and quantifying the solution after 10 minutes. Figure S16 shows the
th
comparison of enzyme activity after 10 minutes on the gel on the first day to the 30 day. Although
less active, it was very interesting to see both enzymes still being able to catalyze both enzymatic
reactions over this long time period.
Figure 6. Recycling experiments with both enzymes entrapped in L and D amino acid hydrogels. a)
MA-
of the gel each day and product was quantified at absorbance 420 nm. b) MA-
with HRP enzyme entrapped. A fresh substrate solution was placed on top of the gel each day and
product was quantified at absorbance 420 nm. c) MA- -Phe-OH hydrogel with amylase enzyme
entrapped. Fresh substrate solution was placed on top of the gel each day and product was quantified
at absorbance 540 nm. d) MA- -Phe-OH hydrogel with an amylase enzyme entrapped. Fresh
L-Phe-OH hydrogel with an HRP enzyme entrapped. Fresh substrate solution was placed on top
D-Phe-OH hydrogel
L
D
substrate solution was placed on top of the gel each day and the product was quantified at absorbance
540 nm.

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3. Discussion
In this study two amino-acid hydrogels were synthesized, characterized, and tested for their
ability to serve as a matrix for enzyme entrapment. Firstly, these hydrogels showed exceptional
stability on their own. They are able to remain in hydrogel state for over a year if no external factors
are involved which itself proves to be a remarkable biomaterial. The hydrogels exhibit thixotropic
properties where upon addition and removal of strain, the gel material breaks and reforms the
hydrogel state. They also have optimal three-dimensional fiber morphology where the nanofibers
and amphiphilic surfaces in the hydrogels allow rapid mass transfer as well as an optimal network
to keep the enzyme entrapped [26]. In addition, the materials show the interesting self-assembling
properties of Phe and the presence of both enantiomers on CD spectroscopy. Phe was found to form
an ordered structure exhibiting a distinct ultra-narrow peak around 220–230 nm, which attributes to
the formation of Phe-based chiral structures that correspond to structural aggregates [40]. These
materials can be tailored by changing the concentration of the gel: A more solid-like material is
formed when the hydrogel concentration is 5% w/v where the material is stiff and less viscous while
a viscous, malleable material is formed when the hydrogel is at its MGC. A working concentration of
1% was chosen for all studies as there is a 30-minute wait period after heating the solution before
reaching a hydrogel state. This provides optimal time to add in the enzyme solution without
deactivating it, thereby allowing the gel to further cool afterwards into a hydrogel with entrapped
enzyme.
Both hydrogels proved to be an optimal matrix for enzyme entrapment as enzymes remained
active in the hydrogel, meaning the enzyme did not completely deactivate and can catalyze the
biochemical reaction while it was entrapped in the gel. To test the ability of the hydrogels to act as a
suitable matrix, the initial activity of both enzymes in the gels were quantified using UV spectroscopy
and their respective extinction coefficients. As time increases with the substrate solution on the gel,
an increase in product formation is observed as there was more time for substrate diffusion and
interaction with the entrapped enzyme. It is also observed that the enzymes do not prefer one
enantiomer over the other, as a very similar activity and quantification results were concluded for
both L and D hydrogels. Enzyme leaching was also explored, and it was found that there was minimal
leakage of the enzyme into the solution on top of the hydrogel. The reported activity using the
enzyme that leached out of the gel into the buffer solution was substantially lower compared to the
activity of the enzyme entrapped in the gel. This reinstates the idea that the interactions between the
amino acid conjugates and the enzyme help to keep the enzyme entrapped, meaning the enzyme
stayed within the hydrogel matrix. In addition, preliminary studies show that the same amount of
enzyme catalyst entrapped in the gel can catalyze fresh substrate over multiple days, allowing the
biocatalyst to be reused, which is beneficial in industrial biotransformations. Future studies involving
testing the kinetics and comparing multiple entrapped enzymes are currently ongoing.
A large number of enzymes trapped within sol–gel matrices show that they usually retain their
catalytic activity while being protected against degradation [49]. Examples of entrapped enzymes
were reported to exhibit better stability, activity, and longer lifetimes than free enzymes, meaning the
enzyme’s ability to catalyze the chemical reaction lasts longer, and converts more product while not
deactivating from external factors such as heat. For example, Xu demonstrated that hydrogelation
provides a useful approach for immobilizing thermally unstable enzymes to significantly enhance
their activity and stability (including thermal and operative) of the enzymes [29]. Compared to
reported examples of enzyme entrapment, these amino acid hydrogels have similar properties where
the stability is prolonged and the enzyme is able to be recycled. However, the number of examples
of amino acid and peptide hydrogels as a matrix for entrapment is limited and should be explored
further. Due to the fact that amino acid and peptide hydrogels resemble extracellular matrices, as
reported in literature [4,5], the enzymes are also able to function in a material that is more
biocompatible and biodegradable. In addition, it was reported that unlike the cross-linked random
networks in a conventional polymeric hydrogel, the larger diffused channels and amphiphilic
surfaces in the molecular hydrogel allow rapid mass transfer of the reactant and the product [29]. The
use of amino acid and peptide hydrogels holds great potential to be widely exploited as a matrix for

Molecules 2019, 24, 2884
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enzyme entrapment. By looking at various amino acid and peptide conjugates with different enzymes
and varying concentrations of hydrogels to affect the pore-sizes or entrapment ability for different
sizes of enzymes, this leads to a potential variety of gels that can be tailored for different catalytic
reactions.
4. Materials and Methods
Methyl esters of amino acids were purchased form Advanced ChemTech, Louisville, KY, US.
Myristic acid, amylase, pyrogallol, starch, and horseradish peroxidase were purchased from Sigma
Aldrich, Mississauga, ON, Canada. Chemicals were used without further purification.
Both
compounds
were
synthesized
using
the
standard
1-ethyl3-(3’-
dimethylaminopropyl)carbodiimide (EDC)/1-hydroxybenzotriazole (HOBt) method. Myrsitic acid (8
mmol) was dissolved in DCM and HOBt (4 mmol, 0.678 g) and EDC (4 mmol, 0.767 g) were added to
the reaction mixture at 0 °C and the reaction mixture was stirred for 30 minutes in ice. Five mmol of
L- or D-Phe-OH was added to the solution and was stirred for 24 hours at r.t. Afterwards, a standard
aqueous workup was done consisting of sequential washing with aqueous 10% citric acid solution,
saturated aqueous NaHCO3, and saturated brine solution. This was followed by collection and drying
of the organic layer. The crude material was then purified by column chromatography on silica gel
using hexane/ethyl acetate solvent system to afford the pure product as a white solid. The compound
was then hydrolyzed to remove the OMe protecting group by adding 1.3 mmol (0.5 g) of myristyl-
phenylalanine-OMe to a round bottom flask with 16 mmol (0.65 g) of NaOH along with 20 mL of 1,4-
dioxane and 20 mL of dH2O. The reaction was stirred at room temperature overnight. After the
stirring, concentrated HCl was added to precipitate the solution and acidify it to pH 2–3. This was
followed by two washes with ethyl acetate. The ethyl acetate layers were collected, dried with
anhydrous sodium sulfate, filtered, and evaporated under pressure. This resulted in the pure white
powder compound used for gelation.
1
MA-L-Phe-OH H-NMR (500 MHz, [D] chloroform): 12.70 (s, 1H, OH), 7.32 (m, 3H; H-Phe), 7.20
(m, 2H; H-Phe), 5.82 (d, 1H; amide NH), 4.86 (td, 1H; CHα), 3.28 (dd, 1H; CH2β), 3.16 (dd, 1H; CH2β),
13
2.20 (td, 2H; (CH2), 1.58 (td, 2H; (CH2)), 1.27 (m, 20H; (CH2)), 0.90 (t, 9H; (CH3)); C-NMR (125 MHz,
[D6] dimethyl sulfoxide): 173.17 (1C; COOH), 172.09 (1C; C=O) 137.75 (1C; benzene), 129.01 (2C;
benzene), 128.00 (2C; benzene), 126.22 (1C; benzene), 53.19 (1C; α-CH), 36.75 (1C; β-CH2), 22.09–35.06
-1
-1
-1
(11C; CH2)), 13.90 (1C; (CH3)); FTIR: 3300 cm (amide N-H), 1705 cm (ester C=O), cm (amide C=O);
ESI-MS: m/z: 375.2773 [M]; Found m/z 376.2851 ([C23 H37 N O3]+H)+; elemental analysis for
C23H37NO3: C 73.56, H 9.93; N 3.73; found: C 73.85, H 9.46, N 3.67.
1
MA-D-Phe-OH H-NMR (500 MHz, [D] chloroform): 12.71 (s, 1H, OH), 7.32 (m, 3H; H-Phe), 7.19
(m, 2H; H-Phe), 5.85 (d, 1H; amide NH), 4.87 (td, 1H; CHα), 3.28 (dd, 1H; CH2β), 3.16 (dd, 1H; CH2β),
13
2.20 (td, 2H; (CH2), 1.57 (td, 2H; (CH2)), 1.28 (m, 20H; (CH2)), 0.90 (t, 9H; (CH3)); C-NMR (125 MHz,
[D6] dimethyl sulfoxide): 173.18 (1C; COOH), 172.09 (1C; C=O) 137.75 (1C; benzene), 129.01 (2C;
benzene), 128.01 (2C; benzene), 126.23 (1C; benzene), 53.19 (1C; α-CH), 36.75 (1C; β-CH2), 22.09–35.06
-1
-1
-1
(11C; CH2)), 13.90 (1C; (CH3)); FTIR: 3278 cm (amide N-H), 1722 cm (ester C=O), 1649 cm (amide
C=O); ESI-MS: m/z: 375.2773 [M]; Found m/z 376.2846 ([C23 H37 N O3]+H)+; elemental analysis for
C23H37NO3: C 73.56, H 9.93; N 3.73; found: C 73.75, H 9.82, N 3.83.
4.1. Nuclear Magnetic Resonance (NMR) Spectroscoopy
Compounds were characterized using NMR spectroscopy on a Bruker 500 MHz spectrometer.
The sample was dissolved in [D6] dimethyl-sulfoxide at a concentration of 2 mm. Spectra are shown
in the Supporting Information.
4.2. Fourier Transform—Infrrared Spectroscopy
-1
FTIR spectra of the compound were recorded in the range 3400–400 cm using a Bruker Alpha
FTIR spectrometer equipped with a diamond for attenuated total reflection. A wet viscous sample

Molecules 2019, 24, 2884
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was prepared by dissolving both compounds in the phosphate buffer, forming the gel, then keeping
the samples under the vacuum to dry.
4.3. Transmission Electron Miscroscopy (TEM)
The morphology of the self-assembled compounds was studied using TEM. The sample was
prepared by dipping our grid into the viscous hydrogel material on a copper TEM grid (300 mesh
size), donated by FORTA Science, coated with Formvar and a carbon film. The grid was allowed to
dry by slow evaporation in air and then further dried in a vacuum for 24 hours. Images were acquired
using a Hitachi 7500 transmission electron microscope. The high contrast of the low-Z self-assembled
material can possibly be due to the thermodynamically stable structure on the Formvar coating.
4.4. Circular Dichroism
Compounds CD spectroscopy was used for determining the chiral molecular arrangement of its
supramolecular assemblies, in gelling (phosphate buffer). CD spectra were recorded between 170 and
600 nm using a Jasco J-810 spectrometer (Easton, MD, USA). Experiments were performed by placing
the 1% gel material into a quartz cuvette of 1 mm path length and measurements were completed in
triplicates.
4.5. Rheology Studies
The viscoelastic properties of the material were studied and frequency sweep rheology
experiments were performed to study the mechanical strength of the gels. Data was acquired using
a Discover Hybrid Rheometer from TA instruments with a 20 mm, 1.997-degree cone, plate Peltier
plate aluminum geometry and were completed in triplicate measurements.
4.6. Enzyme Entrapment, Activity, Leaching, and Quantification Using Ultraviolet (UV) Scpectroscopy
50 μL of a 1 mg/20 mL solution of each enzyme was added into the gel solution upon cooling.
Specifically, 10 mg of each compound was added to a vial with 1 mL of 50 mM monobasic phosphate
buffer at pH 7. This solution was heated using a heat gun until all contents were dissolved and a clear
solution was obtained. Upon cooling a hydrogel forms after roughly 30 minutes at this concentration.
After 15 minutes of leaving the clear solution on the benchtop, where the solution is cooled to the
touch but still in solution form, the enzyme solutions were added in and mixed thoroughly. After the
addition of enzyme solution, we leave the vial on the bench to cool for longer and after roughly 10–
15 minutes the hydrogel with the enzyme inside forms.
For enzyme leaching experiments, 2 mL of buffer was placed on top of the enzyme-entrapped
hydrogels. 50 μL aliquots of buffer were taken out at various times points and were added to fresh
substrate solution to see if the buffer aliquots could catalyze a reaction. The substrate solution for the
HRP enzyme consists of 2.6 mL of 50 mM monobasic phosphate buffer pH 7, 0.16 mL of a 30% H2O2
solution, and 0.32 mL of a 50 mg/mL pyrogallol solution. The substrate solution for the amylase
enzyme consists of 1 mL of a 100 mg/10 mL solution of starch, and 1 mL of 50 mM monobasic
phosphate buffer pH 7. If the aliquots of buffer drawn up can catalyze a separate reaction, it is an
indication that the enzyme has leached out of the gel and is just catalyzing the solution on top of the
gel rather than from within the gel. The amount of activity can be traced back to how much enzyme
leaches out. We have shown that there is not nearly as much activity from the aliquot drawn up,
meaning the majority of the catalysis is within the gel.
For quantification of activity, the substrate solutions for each enzyme were placed on top of the
gel with the enzyme entrapped. The substrate solution on the HRP-entrapped gel was measured at
30 seconds, 1 min, 3 min, 5 min, 10 min, 1 hour, and 24 hours on the UV spectrophotometer at 420
nm, and the substrate solution on the amylase-entrapped gel was measured at 1, 3, 5, 10, and 30
minutes on the UV spectrophotometer at 540 nm. The products were then quantified using their
respective extinction coefficients and the beer-lambert law.

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For recycling and stability experiments, enzymes were entrapped in both hydrogels and
substrate solution was placed on top of the gel every day for a week. The substrate solutions were
quantified after 10 minutes on the gel each day to observe the activity of the enzyme over time. In
addition, the enzyme was placed in a hydrogel and was left on the bench top at room temperature
(about 21 degrees) for one month, after this time substrate solution was placed on top to see if the
enzyme was still active by measuring the solution on top of the gel using the UV spectrophotometer
at each respective wavelength and extinction coefficient to quantify.
5. Conclusions
In this study, two hydrogels were synthesized, characterized, and explored as an optimal matrix
for enzyme entrapment. It was found that these hydrogels are extremely stable over time, retaining
their viscoelasticity properties for one year. Besides the different chiralities, both compounds
exhibited similar morphologies, strength, and abilities to entrap enzymes while retaining activity.
Both compounds were able to entrap two enzymes—horseradish peroxidase and α-amylase—and
the quantification of the product over time was performed. The ability of the enzyme to retain activity
for long periods of time and for the enzyme biocatalyst to be reused after multiple days on a new
substrate solution was also shown. These materials have great potential and show promise to
advance the use of enzymes as biocatalysts on industrial scales for greener, more selective, and
cheaper chemical production.
1
Supplementary Materials: Figure S1. H-NMR spectrum of MA-L-Phe-OH. Figure S2. 13C-NMR spectrum of
MA-L-Phe-OH. Figure S3. ESI-MS spectrum of MA-L-Phe-OH. Figure S4. FTIR spectrum of MA-L-Phe-OH.
Figure S5. 1H-NMR spectrum of MA-D-Phe-OH. Figure S6. 13C-NMR spectrum of MA-D-Phe-OH. Figure S7. ESI-
MS spectrum of MA-D-Phe-OH. Figure S8. FTIR spectrum of MA-D-Phe-OH. Figure S9. Histograms of TEM fiber
thickness for a) MA-L-Phe-OH and b) MA-D-Phe-OH. Figure S10. Additional TEM images of both hydrogels with
entrapped enzymes. a–c) M-L-Phe-OH hydrogel with HRP enzyme entrapped. Scale bar is 2, 2, and 1
m
respectively. d–f) M-L-Phe-OH hydrogel with amylase enzyme entrapped. Scale bar is 2, 1, and 1 m respectively.
g–i) M-D-Phe-OH hydrogel with HRP enzyme entrapped. Scale bar is 2 m for all. j–l) M-D-Phe-OH hydrogel
with amylase enzyme entrapped. Scale bar is 2 m for all. Figure S11. Rheology of hydrogels with entrapped
enzyme. a) MA-L-Phe-OH + HRP. b) MA-D-Phe-OH + HRP. c) MA-L-Phe-OH + amylase. d) MA-D-Phe-OH +
amylase. Figure S12. Time versus absorbance plot of purpurogallin product formation at 420 nm using the HRP
enzyme in solution. Due to the addition of color reagent needed to detect the maltose product, the color change
to red indicates the active free amylase enzyme in solution. Figure S13. Standard concentration versus
absorbance spectrum maltose with color reagent for determination of extinction coefficient. Figure S14. Enzyme
leaching experiment. Here the activity of the enzyme in the gel with substrate solution placed on the gel is
compared to the activity of the catalytic reactions using the buffer solution on the enzyme in gel as the enzyme
source. Here it is determined whether the activity comes from the enzyme leaching out the gel and catalyzing
the reaction in the solution on top of the gel or the catalytic reaction occurring through the gel. a) Activity
comparison of HRP enzyme in MA-L-Phe-OH hydrogel with substrate solution on top (dark blue) versus activity
of catalytic reaction using the enzyme leached out of gel in solely buffer on top of the gel as the enzyme source
(light blue). b) Activity comparison of HRP enzyme in MA-D-Phe-OH hydrogel with substrate solution on top
(dark blue) versus activity of catalytic reaction using the enzyme leached out of gel in solely buffer on top of the
gel as the enzyme source (light blue). c) Activity comparison of amylase enzyme in MA-L-Phe-OH hydrogel with
substrate solution on top (dark blue) versus activity of catalytic reaction using the enzyme leached out of gel in
solely buffer on top of the gel as the enzyme source (light blue). d) Activity comparison of amylase enzyme in
MA-D-Phe-OH hydrogel with substrate solution on top (dark blue) versus activity of catalytic reaction using the
enzyme leached out of gel in solely buffer on top of the gel as the enzyme source (light blue). Figure S15. a)
Image of two L followed by the last two vials of D hydrogels, the first and third vial containing the HRP enzyme
and one does not. The substrate solution is placed on top all four vials while the HRP enzyme was added to the
second and fourth solution (the vials without HRP entrapped). A color change in the hydrogels is observed for
the gels with enzyme entrapped as substrate solution diffuses in and the colored product is formed. For the
hydrogels that do not have the enzyme entrapped in the gel no color change is seen in the gel, only in the
substrate solution. b) Closer view of the vials with and without the enzyme entrapped. Figure S16. Stability test
of enzymes in the gel for one month. a) Comparison of activity of HRP enzyme in MA-L-Phe-OH hydrogel on
day 1 to day 30. b) Comparison of activity of HRP enzyme in MA-D-Phe-OH hydrogel on day 1 to day 30. c)

Molecules 2019, 24, 2884
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Comparison of activity of amylase enzyme in MA-L-Phe-OH hydrogel on day 1 to day 30. d) Comparison of
activity of amylase enzyme in MA-D-Phe-OH hydrogel on day 1 to day 30.
Author Contributions: conceptualization, N.F., H.B.K; methodology, N.F, formal analysis, N.F.; investigation,
N.F.; T.S., R.R, resources, H.B.K., writing—original draft preparation, N.F; writing—review and editing, N.F,
H.B.K; visualization, N.F; supervision, H.B.K; Authorship must be limited to those who have contributed
substantially to the work reported.
Funding: This research received no external funding.
Acknowledgments: We appreciate and acknowledge the funding from the National Sciences and Engineering
Research Council (NSERC) and the University of Toronto. We would also like to thank our TRACES lab manager
Tony Adamo for all of his help and support throughout these studies and Durga Acharya for helping acquire
the TEM images.
Conflicts of Interest: The authors declare no conflict of interest.
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Sample Availability: Samples of the compounds; MA-F(L/D)-OH are available from the authors upon request.
© 2019 by the authors Licensee MDPI, Basel, Switzerland. This article is an open access
article distributed under the terms and conditions of the Creative Commons Attribution
(CC BY) license (http://creativecommons.org/licenses/by/4.0/).