Lipase-Powered Mesoporous Silica Nanomotors for Triglyceride Degradation

Article abstract of DOI:10.1002/anie.201900697

We report lipase-based nanomotors that are capable of enhanced Brownian motion over long periods of time in triglyceride solution and of degrading triglyceride droplets that mimic “blood lipids”. We achieved about 40 min of enhanced diffusion of lipase-modified mesoporous silica nanoparticles (MSNPs) through a biocatalytic reaction between lipase and its corresponding water-soluble oil substrate (triacetin) as fuel, which resulted in an enhanced diffusion coefficient (ca. 50 % increase) at low triacetin concentration (<10 mm). Lipase not only serves as the power engine but also as a highly efficient cleaner for the triglyceride droplets (e.g., tributyrin) in PBS solution, which could yield potential biomedical applications, for example, for dealing with diseases related to the accumulation of triglycerides, or for environmental remediation, for example, for the degradation of oil spills.

Full text of DOI:10.1002/anie.201900697

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Accepted Article
Title: Lipase-powered mesoporous silica nanomotors for triglyceride
degradation
Authors: Lei Wang, Ana Cândida Hortelão, Xin Huang, and Samuel
Sanchez
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To be cited as: Angew. Chem. Int. Ed. 10.1002/anie.201900697
Angew. Chem. 10.1002/ange.201900697
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10.1002/anie.201900697
Angewandte Chemie International Edition
COMMUNICATION
Lipase-powered mesoporous silica nanomotors for triglyceride
degradation
Lei Wang[a,b]+, Ana C. Hortelão[a]+, Xin Huang[b] and Samuel Sanchez[a,c]*
drugs to cells in vitro, as well as for the fabrication of antibody-
modified urease nanomotors for efficient bladder cancer
spheroids.[15] Apart from spherical particles, enzymes have been
reported to power various other structures, such as
supramolecular stomatocytes,[16] polymersomes,[17] carbon
nanotubes,[18] silica nanotubes,[19] PEDOT/Au microtubes,[20]
polypyrrole/Au nanorods,[21] silk fibroin,[22] and macroscale carbon
fibers.[23] Despite significant advances in the field of motors, only
a few micro/nanomotors demonstrated the motion in oily
solution[24], however, enzymatic motors capable of navigating in
oil solution or at the oil-water interface are still rarely explored.
We demonstrate that lipase-powered nanomotors (LNMs), with
stochastic binding of lipase on mesoporous silica nanoparticles
(MSNPs), can swim in dissolvable triglyceride solution, thus
improving the degradation efficiency of slightly-dissolvable
triglyceride. Lipase (triacylglycerol ester hydrolases from Candida
rugosa, EC 3.1.1.3) was chosen for the experiment since lipase,
per se, can act as catalyst for the decomposition of triglyceride
substrates, both dissolvable (e.g., triacetin with solubility of 61
mg/mL) and slightly-dissolvable (e.g., tributyrin with solubility of
0.133 mg/mL). It offers two functionalities: (i) it acts as the power
engine with triacetin as fuel, and (ii) it takes on the role of active
cleaner for the tributyrin droplets. In addition to enhancing the
motion of LNMs, the fuel of triacetin is biocompatible (an FDA
approved food additive[25]) and thus suitable for biological and
environmental applications. Furthermore, LNMs present a high
efficiency in triglyceride degradation compared with free lipase
and other enzymatic motors (See Figure 4).
Abstract: We report lipase-based nanomotors, capable of enhanced
Brownian motion over long periods in triglyceride solution and of
degrading triglyceride droplets that mimic ‘blood lipids’. We achieved
about 40 minutes of enhanced diffusion of lipase modified
mesoporous silica nanoparticles (MSNPs) through a biocatalytic
reaction between lipase and its corresponding water-soluble oil
substrate (triacetin) as fuel, which resulted in an enhanced diffusion
coefficient (i.e., ~50% increase) at low triacetin concentration (e.g., <
10 mM). Lipase does not only serve as the power engine but also as
a highly efficient cleaner for the triglyceride droplets (e.g., tributyrin)
in PBS solution, which could yield potential biomedical applications,
e.g., for dealing with diseases related to the accumulation of
triglycerides, or for environmental remediation, e.g., degradation of oil
spills.
Synthetic micro/nanomotors are active colloids that self-propel in
fluids while performing complex tasks such as cargo pick-ups and
release,[1]
sensing,[2]
biomedical
applications,[3]
and
environmental remediation.[4] There are several methods to
trigger the propulsion of micro/nanomotors, including through
catalytic reactions,[5] electric fields,[6] as well as ultrasonic,[7]
magnetic,[8] and light excitation.[9] Particularly for in vivo
applications, it is necessary to explore biocompatible and
biomimetic designs, capable of actuating in biological systems
without side effects. Therefore, enzymes that carry out energy
conversion in biological systems by using existing fuel sources,
and without requiring an external power source are good
candidates to address these concerns.[10] Sen et al. provided
experimental evidence of enhanced single-enzyme diffusion
based on substrate turnover.[11] They also demonstrated
molecular chemotaxis of urease and catalase towards a gradient
of increasing substrate concentrations,[11b] as well as the
fabrication of enzymatic motors.[12] Our group extended this field
by modifying the Janus mesoporous silica micro/nanoparticles
with different enzymes.[13] With urease, the hollow Janus silica
micromotors achieved a directional propulsion with a high degree
of controllability, either through the addition of salts or in the
presence of a magnetic field.[13b] In addition, we reported how the
stochastic binding of enzymes to the surface of the particles with
a molecular asymmetry in enzyme distribution results in an
asymmetric generation of forces which in turn leads to self-
propulsion.[14] This technique was also adopted for the fabrication
of nanomotors based on mesoporous silica nanoparticles which
yielded a more efficient delivery and release system of anticancer
[a]
[b]
Dr. L. Wang, A. C. Hortelão, Prof. Dr. S. Sanchez,
Institute for Bioengineering of Catalonia (IBEC), The Barcelona
Institute of Science and Technology (BIST), Baldiri i Reixac 10-
12, 08028 Barcelona, Spain
E-mail: ssanchez@ibecbarcelona.eu
Dr. L. Wang, Prof. Dr. X. Huang,
MIIT Key Laboratory of Critical Materials Technology for New
Energy Conversion and Storage, School of Chemistry and Chemical
Engineering, Harbin Institute of Technology, Harbin 150001, China
Prof. Dr. S. Sanchez,
Institució Catalana de Recerca i Estudis Avançats (ICREA), Pg.
Lluís Companys 23, 08010 Barcelona, Spain
These two authors contributed equally to this work.
Supporting information for this article is given via a link at the end of
the document.
Figure 1. (a) Schematic representation of the functionalization strategy for the
preparation of the lipase-based nanomotors whose motion was triggered by
catalytic reaction with triacetin. (b) SEM and (c) TEM microscopy images of
MSNPs modified lipase, with insets showing magnifications. Scale bars are: (b)
1 µm (100 nm in the inset), and (c) 500 nm (20 nm in the inset). (d) Evolution of
surface charge evolution after modifying the MSNP surfaces with amine
functional groups, followed by lipase. All measurements were carried out in the
same PBS (phosphate-buffered saline, pH 7.4) solution. (e-g) Confocal
fluorescence (FL) images of lipase-modified MSNPs, scale bars are 5 µm. The
red FL is due to Rhodamine B (RB) loading of the MSNPs, while the green FL
is generated by the Krypton-modified lipase. (g) The corresponding merged
image.
[c]
+
LNMs were fabricated using MSNPs as substrate to immobilize
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10.1002/anie.201900697
Angewandte Chemie International Edition
COMMUNICATION
lipase. MSNPs are biocompatible and their porous structure is
useful for drug loading.[26] The surface of the MSNPs was
modified chemically using glutaraldehyde as crosslinker between
the amino groups of lipase and MSNPs, respectively (Figure 1a),
resulting in stable catalytic activity.[27] MSNPs were characterized
by transmission electron microscopy (TEM) and scanning
electron microscopy (SEM) (Figure S1a-b). They consisted of
uniform spheres (Figure 1b,c) with a porous structure (Figure S1a
inset) and an average diameter of 431.01 ± 1.99 nm (N = 100,
average diameter ± standard error of the mean (SE), cf., Figure
S2). They showed no aggregation which was also confirmed
through dynamic light scattering (DLS). After each modification
step, there was only a single population distribution in the
hydrodynamic radius plots (Figure S3a), indicating that MSNPs
were still well dispersed. The increase in hydrodynamic radius
and broader peak can be attributed to the amination and lipase
immobilization (Figure S3a). Furthermore, the change in surface
charge confirmed each modification step (Figure 1d): the as-
synthesized MSNPs showed a negative surface charge of -16.43
± 1.36 mV, which increased to 22.57 ± 1.79 mV after amination.
Lipase immobilisation in PBS solution (phosphate-buffered saline,
pH 7.4) caused the surface charge to drop to -7.68 ± 1.21 mV ,
which can be attributed to the isoelectric point of lipase of about
pH 5.6~5.8.[28] Additionally, protein immobilization was
characterized via fluorescent staining (Figure 1e to g). Confocal
FL microscopy images showed the MSNPs loaded with
Rhodamine B (RB) (Figure 1e) and stained with Krypton (Figure
1f). Immobilization of lipase on the surface of MSNPs was also
confirmed (Figure 1g). Those results were further confirmed by
the presence of a peak at 280 nm in UV-spectra of lipase-modified
MSNPs (Figure S3b).[29]
PBS solution, we selected triacetin as substrate due to its
solubility in PBS. Since the catalytic reaction of triacetin generates
acetic acid (Equation 1 in SI), litmus (10 mg/mL) was added as
pH indicator to the triacetin (10 mM)-containing PBS solution (pH
7.4, 10 mM) to further prove catalytic activity of the LNMs. This
was followed by the collection of UV spectra. Initially, i.e., prior to
the addition of the LNMs, the triacetin PBS solution showed a blue
colour (absorption at 580 nm, Figure S4b blue curve), which
changed to pink within about 3h after adding the LNMs
(absorption at 500 nm, Figure S4b red curve). This further
confirms that lipase maintained its activity after being immobilized
on MSNPs. We tested different concentrations of triacetin as fuel
for the lipase-modified MSNPs. The mean squared displacement
(MSD) (Figure 2a), calculated from the tracked trajectories (Figure
2a, inset), increased linearly with time and fuel concentration,
indicating a fuel concentration-dependent motility.[30] The effective
diffusion coefficient (De) was calculated from the MSD per time
interval (Δt), as: De = 0.25MSD/Δt. During the studied time range,
De of lipase-modified MSNPs was 0.72 ± 0.03 μm2/s in PBS
solution without adding fuel, increasing to 0.94 ± 0.05, 1.08 ± 0.05,
and 1.13 ± 0.04 μm2/s (Figure 2b) when adding 1.0, 10 and 100
mM of fuel, respectively. This illustrates the increase of De with
triacetin concentration. Since De increased only marginally (by
4.6%) when adding 100 mM instead of 10 mM of fuel, we used 10
mM for all subsequent experiments. When adding 10 mM of fuel,
LNMs were capable of a sustained motion for about 40 min
(Figure 2c). After 40 min, De dropped again to the control value
(sample without added fuel) (Figure 2d).
Figure 2. a) Representative trajectories (inset) of LNMs with different triacetin
concentrations of 0 mM (black), 1 mM (red), 10 mM (green), and 100 mM (blue)
and corresponding mean-squared displacements (MSD) (n ≥ 20, error bars
represent SE); b) effective diffusion coefficients obtained by analysing the MSD
for different triacetin concentrations; c) representative trajectories of LNMs at
different times with sufficient fuel; d) diffusion coefficients of LNMs as a function
of time, with 100 mM of triacetin. Control experiment is LNMs in PBS solution
without fuel. *p<0.05 when compared to the control group.
Figure 3. Representative trajectories of lipase-motors surrounding tributyrin
droplets with (a) 0 and (b) 10 mM of added triacetin. Scale bars are 5 µm. (c)
MSD of the LNMs with 0 and 10 mM triacetin (n ≥ 16, error bars represent the
SE). (d) Effective diffusion coefficients obtained from the MSD at different
triacetin concentrations (with tributyrin droplets present in the solution). (e-f) FL
microscopy images showing the interaction between the droplets (red FL from
Nile Red in tributyrin oil phase) and motors (green FL from FITC loaded in
MSNPs) at different times.
Enzyme assays were carried out to confirm that lipase remained
active after its immobilization onto the MSNPs. In the studied
motor concentration range of 0.1 to 1.0 mg mL-1, the active
milliunits of lipase linearly increased with motor concentration
(Figure S4a), confirming the catalytic activity of lipase-modified
MSNPs. To investigate the motility of lipase-modified MSNPs in
To explore the ability of LNMs to degrade triglyceride, tributyrin
was selected as removal target since it is slightly dissolvable in
PBS solution, thus resembling ‘blood lipid’. When LNMs were
added to a PBS solution containing tributyrin droplets in
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10.1002/anie.201900697
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COMMUNICATION
suspension (Figure S6), LNMs exhibited typical Brownian motion
with De = 0.78±0.043 µm2/s (Figure 3a-d). This suggests that low
tributyrin concentrations (0.133 mg/mL, saturated concentration)
do no induce enhanced Brownian diffusion of LNMs. After adding
fuel (triacetin, 10 mM), the LNMs moved more rapidly resulting in
De = 1.096±0.085 µm2/s (Figure 3b-d). When the LNMs happen
to reach the surface of the oil droplets, they are confined at the
oil-water interface due to the inherent amphiphility of lipase,[31] but
still maintain a diffusion coefficient of De = 0.174±0.038 µm2/s
(Figure S7), that is significantly smaller than that of Brownian
motion in bulk solution. The confinement of LNMs to the oil
droplets’ surface is shown in Figures 3e-f and S8, where the
interfacial catalysis between lipase and tributyrin occurred.
Therefore, tributyrin droplets were degraded (≤20 µm in diameter)
within 1h.
of the surrounding solution was simultaneously analysed via
individual channel using Image J software (RGB intensity, cf.,
Figure 4b). Initially, the diameter of a representative tributyrin
droplet was about 11.36 µm (Figure 4d) and the solution exhibited
a blueish colour (blue intensity (BI): 231.6; red intensity (RI): 160)
with a pH value of 7.4 (Figure 4b, t=0 min). After 10 minutes, the
diameter of the droplet had decreased to 9.65 µm and the
surrounding solution turned purple (Figure 4a-b, BI:203.5 and
RI:194.1, pH 4.5). As time progressed, the droplet size continued
to decrease, while the surrounding solution turned increasingly
red (decreasing BI and increasing RI values). Toward the end of
the experiment (50 min), the tributyrin droplet was degraded and
the solution had changed to pink (BI:160.0; RI:207.7), due to the
continuous release of butyric acid into the surrounding solution.
This represents a visible dynamic demonstration of the tributyrin
degradation process (from 0% to 100%).
To further demonstrate the advantages of LNMs for the
degradation of tributyrin, different control experiments were
conducted (Figure 4e-f, S9) including: bare MSNPs, triacetin,
urease-MSNP nanomotors, urease-MSNP nanomotors with the
fuel of urea (20 mM), free lipase, free lipase with triacetin, LNMs
without fuel, and LNMs with triacetin as fuel (10 mM). Urease-
nanomotors were chosen as controls because we needed to
determine whether enhanced diffusion can improve the
degradation. With bare MSNPs there was only 5.2% degradation
of the droplets after 1h. This is mainly due to the low tributyrin
solubility, indicating that bare MSNPs have almost no effect on
tributyrin droplets. With pure triacetin (10 mM), urease-MSNP
motors with urea (20 mM), and urease-MSNP motors without urea
yielded degradations of 1%, 6.5%, and 6.7%, respectively. We
can thus conclude that urease-MSNP nanomotors, despite
containing enzymes on the surface and presenting enhanced
diffusion, cannot degrade tributyrin droplets. Additionally, we also
compared the affinities of urease motors and LNMs for targeting
the tributyrin droplets. The same concentration of MSNPs were
later separated into two aliquots and loaded with fluorescein
isothiocyanate (FITC), followed by the corresponding modification
with urease and lipase, respectively. After these two types of
FITC-loaded nanomotors were added into the solution containing
tributyrin droplets, we calculated the FL intensity on the tributyrin
droplet area (details in SI, Figures S10-11). Initially, both types of
motors distributed randomly in the solution (Figure S10) with a
similar FL intensity within the droplet area. Within 1h, the FL
intensity surrounding the droplet area increased dramatically in
the presence of fuel for the LNMs (Figure S11), confirming that
LNMs tend to accumulate on the surface of tributyrin droplets
which leads to their highly efficient degradation. Besides, with free
lipase (2 µg/mL) the droplet degradation was 48.9%, which
increased to 57.9% after adding triacetin, thus illustrating the
enhanced degradation process in the presence of fuel due to the
enhanced diffusion of free lipase. LNMs (50 µg/mL) with 2 µg/mL
lipase (measured through bicinchoninic acid assay (BCA), details
in SI) can lead to a 78.8% degradation, i.e., higher than with free
lipase. This is because free lipase tends to form bimolecular
aggregates that partially block the lipase active centres, which in
turn results in a less active form.[32] The immobilization can be
beneficial for cleaving these lipase dimers, thus improving their
efficiency. Notably, more than 98% of the droplets can be
degraded through the addition of triacetin, which demonstrates
how lipase anchored to MSNPs increases the efficient propulsion
and diffusion towards the target.
Figure 4. Demonstrating the dynamic shrinking process of tributyrin droplets
with lipase-motors at different times. (a) The colour change of the surrounding
medium is caused by the litmus indicating the continuous generation of butyric
acid. Round dashed circles indicate the initial size of the tributyrin droplets. The
scale bar is 5 µm. (b) Corresponding RGB intensity and pH value in the
surrounding solution. (c) Size measurement of the tributyrin droplets according
to the white dash line in (a). (d) Degradation percentage of tributyrin droplets
calculated from their diameters (see inset); the chemical equation shows the
catalytic decomposition of tributyrin. (e) Optical microscopy of the degradation
processes of tributyrin droplets under different conditions. (f) Corresponding
histogram plot to (e), showing the degradation percentage of tributyrin droplets
under different conditions after 1h.
In order to dynamically explore the degradation process, we focus
on a single representative tributyrin droplet. Litmus was used to
illustrate the dynamic colour change during the degradation of
tributyrin by LNMs which produces butyric acid (Figure 4a,
Equation 2 in SI). While the fuel-induced catalytic reaction can
trigger an immediate local pH change, the colour change may be
delayed by up to three hours due to the low fuel concentration (10
mM). However, tributyrin degradation significantly accelerated the
acidification, producing a colour change in just 30 min. The colour
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10.1002/anie.201900697
Angewandte Chemie International Edition
COMMUNICATION
10, 839-844; d) J. Wang, Z. Xiong, J. Zheng, X. Zhan, J. Tang,
Accounts Chem. Res. 2018, 51, 1957-1965.
In conclusion, we have demonstrated that the lipase-based
catalytic reaction can be utilized to provide the enhanced motility
of MSNPs in triglyceride-containing PBS solutions. Compared to
free lipase and urease nanomotors, we showed that active lipase-
nanomotors can accelerate the degradation process of tributyrin
droplets to a very high degree (~98% within 50 min). These results
hint towards possible new applications of lipase-powered
nanomotors, in biomedicine for high-triglyceride related diseases,
or in environmental remediation for oil removal. However, it is
necessary to understand the stability and activity of such motors
in oil-polluted waters, as well as the interaction of these motors
with adipocytes. Future work should be dedicated to improving
the motion control, as well as their performance in biological fluids,
like serum or blood.
[10] a) T. Patiño, X. Arqué, R. Mestre, L. Palacios, S. Sánchez,
Accounts Chem. Res. 2018, 51, 2662-2671; b) X. Zhao, K.
Gentile, F. Mohajerani, A. Sen, Accounts Chem. Res. 2018,
51, 2373-2381; c) X. Ma, A. C. Hortelão, T. Patiño, S. Sánchez,
ACS nano 2016, 10, 9111-9122.
[11] a) H. S. Muddana, S. Sengupta, T. E. Mallouk, A. Sen, P. J.
Butler, J. Am. Chem. Soc. 2010, 132, 2110-2111; b) S.
Sengupta, K. K. Dey, H. S. Muddana, T. Tabouillot, M. E. Ibele,
P. J. Butler, A. Sen, J. Am. Chem. Soc. 2013, 135, 1406-1414.
[12] K. K. Dey, X. Zhao, B. M. Tansi, W. J. Méndez-Ortiz, U. M.
Córdova-Figueroa, R. Golestanian, A. Sen, Nano Lett. 2015,
15, 8311-8315.
[13] a) X. Ma, A. Jannasch, U.-R. Albrecht, K. Hahn, A. Miguel-
López, E. Schäffer, S. Sánchez, Nano Lett. 2015, 15, 7043-
7050; b) X. Ma, X. Wang, K. Hahn, S. Sánchez, ACS nano
2016, 10, 3597-3605; c) X. Ma, S. Sánchez, Tetrahedron 2017,
73, 4883-4886.
[14] a) T. Patiño, N. Gracia, X. Arqué, A. Miguel-López, A.
Jannasch, T. Stumpp, E. Schäffer, L. Albertazzi, S. Sánchez,
J. Am. Chem. Soc. 2018, 140, 7896-7903; b) T. Patino, A.
Porchetta, A. Jannasch, A. Lladó, T. Stumpp, E. Schäffer, F.
Ricci, S. Sánchez, Nano Lett., 2019, doi: 10.1021/
acs.nanolett.8b04794.
[15] a) A. C. Hortelão, T. Patiño, A. Perez-Jiménez, À. Blanco,
S. Sánchez, Adv. Funct. Mater. 2018, 28, 1705086; b) A. C.
Hortelão, R. Carrascosa, N. Murillo-Cremaes, T. Patiño, S.
Sánchez, ACS Nano 2018, 13, 429-439.
[16] L. Abdelmohsen, M. Nijemeisland, G. Pawar, G. Janssen, R.
J. Nolte, J. van Hest, D. Wilson, ACS Nano 2016, 10, 2652-
2660.
[17] A. Joseph, C. Contini, D. Cecchin, S. Nyberg, L. Ruiz-Perez,
J. Gaitzsch, G. Fullstone, X. Tian, J. Azizi, J. Preston, G. Volpe,
G. Battaglia, Sci. Adv. 2017, 3, e1700362.
Acknowledgements:
The authors thank Albert Miguel López and Lucas Santiago
Palacios Ruiz for the development of the tracking and extracting
software, respectively. L.W. thanks NSFC (No. 51703043), the
Marie Skłodowska-Curie fellowship (Grant No. 712754) and the
Severo Ochoa programme (Grant SEV-2014-0425 (2015-2019)).
A.C.H. thanks MINECO for the Severo Ochoa fellowship. X. H.
thanks NSFC (No. 21871069). S.S. thanks the Spanish MINECO
for grants CTQ2015-68879-R (MICRODIA) and CTQ2015-72471-
EXP (Enzwim).
Keywords: enzyme-powered nanomotors
•
triglyceride
degradation • oil removal • self-propulsion • MSNPs
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10.1002/anie.201900697
Angewandte Chemie International Edition
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Lipase-based nanomotors capable of
Lei Wang, Ana C. Hortelão, Xin Huang,
and Samuel Sanchez
self-propelling
and
degrading
triglyceride in oil-containing solution for
long periods are reported.
Page No. – Page No.
Lipase-powered mesoporous silica
nanomotors for triglyceride degradation
This article is protected by copyright. All rights reserved.