Enzyme promotes the hydrogelation from a hydrophobic small molecule

Article abstract of DOI:10.1021/ja9042142

(Chemical Equation Presented) We report in this paper that an enzymatic reaction can be used as the sole mechanism for forming supramolecular hydrogels that have good stability in aqueous solutions at room temperature. The gels are two-component hydrogels that are mainly formed by hydrophobic compound 2 and doped with hydrophilic compound 1. They were formed by an enzymatic process that produces hydrophobic 2 in homogeneous modes and assists the formation of three-dimensional networks. We have characterized the morphologies of the gels with scanning electron microscopy and dark-field transmission electron microscopy, collected fluorescence data to monitor the gelation process, and proposed a possible mechanism to explain the formation of the gels.

Full text of DOI:10.1021/ja9042142

Published on Web 07/24/2009
Enzyme Promotes the Hydrogelation from a Hydrophobic Small Molecule
Jie Gao,† Huaimin Wang,† Ling Wang,*,§ Jingyu Wang,† Deling Kong,*,† and Zhimou Yang*,‡
Key Laboratory of BioactiVe Materials, Ministry of Education, Tianjin Key Laboratory of Protein Science,
College of Life Sciences, and College of Pharmacology, Nankai UniVersity, Tianjin 300071, P. R. China
Received May 25, 2009; E-mail: chwling@gmail.com; kongdeling@nankai.edu.cn; chemyzm@gmail.com
In this paper, we report on the use of an enzymatic process to
assist the formation of supramolecular hydrogels from a hydro-
phobic compound. Supramolecular hydrogels,1 composed of three-
dimensional (3D) networks formed by the self-assembly of small
molecules in aqueous solutions, have attracted intense research
interest in recent years. They have been used as a platform for
biosensing,2 a 3D matrix for cell cultures,3 an encapsulation material
for drug release,4 etc. Molecules that can form supramolecular
hydrogels (so-called molecular hydrogelators) are usually am-
phiphilic, such as derivatives of amino acids,5 peptides,6 and
carbohydrates.7 Hydrogels formed by hydrophobic compounds have
never been reported. Since hydrogels formed by hydrophobic
compounds might be more stable than those formed by amphiphilic
compounds, the formation of hydrogels from hydrophobic com-
pounds is desired. In this study, we researched the use of a
phosphatase to convert a hydrophilic precursor to a hydrophobic
compound in a homogeneous mode, thus resulting in the formation
of supramolecular hydrogels with good stability.
Enzymes favor the self-assembly of small molecules, and it has
been proven that enzymatic conversions promote the formation of
more ordered structures in supramolecular hydrogels.8 We envi-
sioned that the enzymatic process, in addition to helping form
ordered structures, may also be the sole mechanism by which
supramolecular hydrogels are formed in some special cases. To
prove this hypothesis, we designed and synthesized 1 as a substrate
of phosphatase. Hydrophobic 2 was generated by treatment with
the enzyme. We thought that gels would be the result of this process
because the enzyme produced 2 in a homogeneous mode, which
promoted the formation of the 3D fiber networks that served as
the matrix of the hydrogels.
II, respectively.9 The gels did not swell and were stable for at least
1 month at pH values from 0 to 9 (e.g., gel II in Figure 1F,G)
below 25 °C. This observation clearly demonstrated that enzymatic
conversion provides a sole mechanism for promoting the formation
of self-assembled systems.
Figure 1. Optical images of 0.5 mL of (A) 1.0 wt % 2 in PBS buffer (pH
8.0) after sonication; (B) 1.0 wt % 1 in PBS buffer (pH 8.0); (C) suspension
formed by the solution in (B) with enzyme (20 units/mL final concentration);
(D) gel I formed by the solution in (B) with 16 units/mL of enzyme; (E)
gel II formed by the solution in (B) with 4 units/mL of enzyme; and gel II
in aqueous solution at (F) pH 0.0 and (G) 9.0.
Scanning electron microscopy (SEM) was used to characterize
the gels. As shown in Figure 2A, large films with widths of
hundreds of micrometers and thicknesses of ∼1.8 µm provided the
matrix of gel I. For gel II (Figure 2B), fibers with widths of 0.3-2
µm (arrows) and thin films with widths of tens to hundreds of
micrometers were entangled with each other to form the 3D
networks. Interestingly, nanoparticles with diameters of 100-800
nm and small pores or dimples adhered to the surfaces of the large
films in gel I (Figure S-2A in the Supporting Information),9
a
The synthesis of 1 was easy and straightforward.9 Treating
L-tyrosine-OMe with Fmoc-OSu in the presence of NaHCO3
resulted in the formation of 2 in high yield (93.4%). First, the
phosphorylation of 2 was conducted with triethyl phosphite and
I2; treatment with TMSBr followed, affording title compound 1 in
85.5% yield. After obtaining both compounds, we tested the gelation
ability of 2 and found that it would not form gels upon pH
adjustment, changes in temperature, or sonication because of its
poor solubility in aqueous solutions [<0.01 mg/mL in PBS buffer
(pH 8.0); Figure 1A]. Next, we tried the enzymatic conversion.
Treating a clear solution of 1 in PBS buffer with high concentrations
of phosphatase (>16 units/mL) resulted in the formation of
suspensions within 1 min (Figure 1C). Interestingly, opaque
hydrogels were formed when lower concentrations of enzyme were
used: final concentrations of 16 and 4 units/mL gave gel I (Figure
1D) and gel II (Figure 1E), respectively. Both gel I and gel II
formed within 10 min, and HPLC results indicated that 64.3 and
62.8% of 1 had been converted at the gelling points of gels I and
phenomenon that was not observed for gel II (Figure S-2B).9 Dark-
field transmission electron microscopy (TEM) images indicated that
the film structures in gels I and II consisted of fibers with widths
of 0.45-2 µm (Figure 2C) and 200-600 nm (Figure 2D),
respectively. The results obtained by SEM and TEM suggest that
higher concentrations of enzyme lead to larger aggregates.
To understand the molecular arrangement of the fluorenyl groups
in the solution and hydrogels, we used fluorescence spectroscopy
to characterize the solution and hydrogels. As shown in Figure 3A,
1 in solution showed an emission peak centered at 310 nm, and
this peak shifted to 323 and 361 nm in both gel I and gel II after
enzymatic conversion, suggesting that the fluorenyl groups in the
two gels overlapped in both antiparallel and parallel manners.10
The higher intensities of the peaks at 323 nm relative to the peaks
at 361 nm indicate that the fluorenyl groups favor the antiparallel
overlap. The peaks at ∼440 nm were from excimer emission of
fluorenyl groups.10 The higher intensities of the peaks in gel I
relative to those in gel II implies that the fluorenyl groups are
stacked more efficiently in gel I, which is consistent with the larger
aggregates in gel I observed by SEM and TEM and the more opaque
appearance of gel I in Figure 1D.
† Ministry of Education.
§ College of Pharmacology.
‡ College of Life Sciences.
9
11286 J. AM. CHEM. SOC. 2009, 131, 11286–11287
10.1021/ja9042142 CCC: $40.75  2009 American Chemical Society

C O M M U N I C A T I O N S
Figure 4. Schematic illustration of enzymatic conversion and proposed
molecular arrangements of 1 and 2 in nanofibers (the antiparallel stacking
of fluorenyl groups is highlighted in green).
Figure 2. SEM images of balanced (A) gel I and (B) gel II and TEM
images of (C) gel I and (D) gel II after 1 day.
of aggregates. The stable hydrogels formed by the enzymes could
be developed into biomaterials that would be useful, for example,
in drug delivery, enzyme immobilizations, and tissue engineering.
We also used fluorescence microscopy to monitor the process
of enzymatic hydrogelation (measured every 10 min after gelation).
As shown in Figure 3B, the intensities of the peaks at 323 and 361
nm in gel II continued to decrease while the intensities of the peaks
at ∼440 nm increased. This result indicates that the enzyme
continued to convert 1 into 2 in the initial stages, thus leading to
more efficient stacking of fluorenyl moieties and more rigid
hydrogels. The conversions stopped after 24 h, as proven by the
HPLC results (Table S-3); the same 1/2 ratio existed in gel II after
24 and 48 h (i.e., there was ∼3.9% 1 in the gel).9
Acknowledgment. This work was supported by the National Out-
standing Youth Fund (30725030), the NSFC (20774050), “973 Projects”
(2007CB914801), and the Tianjin Government (08SYSYTC00200). We
thank Prof. Zhen Xi for help with fluorescence measurements and Mr.
Guo for help with SEM and TEM measurements.
Supporting Information Available: Synthesis and characterization
of 1 and 2, HPLC results, optical images of suspensions with different
ratios of 1 and 2, and SEM images of the two gels and the nanoparticles
in gel I. This material is available free of charge via the Internet at
http://pubs.acs.org.
We propose the following possible explanation of why the enzyme
can help a “nonhydrogelator” form a hydrogel and how the nanofibers
formed by hydrophobic 2 could be stable in aqueous solutions. As
shown in Figure 4, on the basis of the spectroscopic analysis, most of
the fluorenyl groups are stacked in an antiparallel mode, providing
one of the major driving forces for nanofiber formation. We assume
that the nanofibers were mainly formed by 2 and doped with
hydrophilic 1, making the nanofibers stable in aqueous solutions. 1 in
the nanofibers can avoid being hydrolyzed by phosphatase, as
demonstrated by the HPLC results (i.e., 3.9% of 1 was still intact in
gel II after 24 and 48 h).9 Nevertheless, only suspensions could be
obtained from mixtures of different ratios of 1 and 2 upon sonication
or by a heating-cooling cycle,9 further demonstrating that the enzyme
generates hydrophobic 2 in a homogeneous mode, thus assisting the
formation of 3D fiber networks in supramolecular hydrogels.
In summary, we have demonstrated that an enzyme offers a sole
mechanism for generating hydrophobic compounds in homogeneous
modes, thus assisting the formation of 3D fiber networks and
supramolecular hydrogels. This process provides a facial strategy
for generating supramolecular hydrogels from hydrophobic mol-
ecules and offers more candidates for the generation of supramo-
lecular hydrogels. What is more, there is potential for this approach
to be developed into a convenient way to control the morphology
References
(1) Estroff, L. A.; Hamilton, A. D. Chem. ReV. 2004, 104, 1201. Gazit, E.
Chem. Soc. ReV. 2007, 36, 1263. Ulijn, R. V.; Smith, A. M. Chem. Soc.
ReV. 2008, 37, 664. de Loos, M.; Feringa, B. L.; van Esch, J. H. Eur. J.
Org. Chem. 2005, 3615. Vauthey, S.; Santoso, S.; Gong, H. Y.; Watson,
N.; Zhang, S. G. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 5355. Mahler, A.;
Reches, M.; Rechter, M.; Cohen, S.; Gazit, E. AdV. Mater. 2006, 18, 1365.
van Bommel, K. J. C.; van der Pol, C.; Muizebelt, I.; Friggeri, A.; Heeres,
A.; Meetsma, A.; Feringa, B. L.; van Esch, J. Angew. Chem., Int. Ed. 2004,
43, 1663. Yang, Y. G.; Suzuki, M.; Owa, S.; Shirai, H.; Hanabusa, K.
J. Am. Chem. Soc. 2007, 129, 581.
(2) Wada, A.; Tamaru, S.; Ikeda, M.; Hamachi, I. J. Am. Chem. Soc. 2009,
131, 5321. Ojida, A.; Mito-oka, Y.; Sada, K.; Hamachi, I. J. Am. Chem.
Soc. 2004, 126, 2454. Kiyonaka, S.; Sada, K.; Yoshimura, I.; Shinkai, S.;
Kato, N.; Hamachi, I. Nat. Mater. 2004, 3, 58.
(3) Jayawarna, V.; Ali, M.; Jowitt, T. A.; Miller, A. E.; Saiani, A.; Gough,
J. E.; Ulijn, R. V. AdV. Mater. 2006, 18, 611. Storrie, H.; Guler, M. O.;
Abu-Amara, S. N.; Volberg, T.; Rao, M.; Geiger, B.; Stupp, S. I.
Biomaterials 2007, 28, 4608. Hartgerink, J. D.; Beniash, E.; Stupp, S. I.
Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 5133. Holmes, T. C.; de Lacalle,
S.; Su, X.; Liu, G. S.; Rich, A.; Zhang, S. G. Proc. Natl. Acad. Sci. U.S.A.
2000, 97, 6728.
(4) Seo, Y. J.; Bhuniya, S.; Kim, B. H. Chem. Commun. 2007, 1804. Vemula,
P. K.; Li, J.; John, G. J. Am. Chem. Soc. 2006, 128, 8932.
(5) Suzuki, M.; Hanabusa, K. Chem. Soc. ReV. 2009, 38, 967. Yang, Y. G.;
Nakazawa, M.; Suzuki, M.; Shirai, H.; Hanabusa, K. J. Mater. Chem. 2007,
17, 2936.
(6) Zhang, S. G. Nat. Biotechnol. 2003, 21, 1171. Behanna, H. A.; Donners,
J.; Gordon, A. C.; Stupp, S. I. J. Am. Chem. Soc. 2005, 127, 1193.
(7) Komatsu, H.; Matsumoto, S.; Tamaru, S.; Kaneko, K.; Ikeda, M.; Hamachi,
I. J. Am. Chem. Soc. 2009, 131, 5580. Yang, Z. M.; Liang, G. L.; Ma,
M. L.; Abbah, A. S.; Lu, W. W.; Xu, B. Chem. Commun. 2007, 843.
(8) Williams, R. J.; Smith, A. M.; Collins, R.; Hodson, N.; Das, A. K.; Ulijn,
R. V. Nat. Nanotechnol. 2009, 4, 19. Yang, Z. M.; Liang, G. L.; Ma, M. L.;
Gao, Y.; Xu, B. Small 2007, 3, 558. Yang, Z. M.; Liang, G. L.; Wang, L.;
Bing, X. J. Am. Chem. Soc. 2006, 128, 3038.
(9) See the Supporting Information.
(10) Channon, K. J.; Devlin, G. L.; Magennis, S. W.; Finlayson, C. E.; Tickler,
A. K.; Silva, C.; MacPhee, C. E. J. Am. Chem. Soc. 2008, 130, 5487. Smith,
A. M.; Williams, R. J.; Tang, C.; Coppo, P.; Collins, R. F.; Turner, M. L.;
Saiani, A.; Ulijn, R. V. AdV. Mater. 2008, 20, 37. Yang, Z. M.; Gu, H. W.;
Zhang, Y.; Wang, L.; Xu, B. Chem. Commun. 2004, 208.
Figure 3. Emission spectra (λexcitation ) 265 nm) of A) solution of 1, gel
I, and gel II and B) gel II at different time points (every 10 min).
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