Using supramolecular hydrogels to discover the interactions between proteins and molecular nanofibers of small molecules

Article abstract of DOI:10.1039/c2cc33631f

Here we report the first example of the use of supramolecular hydrogels to discover the protein targets of aggregates of small molecules.

Full text of DOI:10.1039/c2cc33631f

View Article Online / Journal Homepage / Table of Contents for this issue
Dynamic Article Links
ChemComm
Cite this: Chem. Commun., 2012, 48, 8404–8406
www.rsc.org/chemcomm
COMMUNICATION
Using supramolecular hydrogels to discover the interactions between
proteins and molecular nanofibers of small moleculesw
Yuan Gao,a Marcus J. C. Long,b Junfeng Shi,a Lizbeth Hedstromab and Bing Xu*a
Received 20th May 2012, Accepted 2nd July 2012
DOI: 10.1039/c2cc33631f
Here we report the first example of the use of supramolecular
hydrogels to discover the protein targets of aggregates of small
molecules.
few examples of the aggregates of small molecules that directly
associate with cellular functions or find applications in biomedicine.
Third, there is no general assay for evaluating the aggregates of
small molecules and their corresponding biological targets.
Since most cells are soft and their interiors are highly viscous and
crowded, we use supramolecular hydrogels to mimic the cellular
environment and have developed supramolecular hydrogel protein
binding assay for discovering the protein targets of the nanofibers of
small molecules. Supramolecular hydrogels (also termed as
‘‘molecular hydrogels’’)10,11 are one kind of heterogeneous,
viscoelastic materials that can immobilize a large amount of water
(up to 99% by weight) by the small molecules (normally less than
1000 Dalton) that self-assemble in water to form nanofibers. Unlike
polymeric hydrogels12 that consist of random networks with small
pore sizes, supramolecular hydrogels consist of only water and the
networks formed by the self-assembly of small molecules. Having
high water content and being viscoelastic, supramolecular
hydrogels closely resemble the crowded cellular environment and
have emerged as a new class of materials for tissue engineering.13
More importantly, Hamachi et al. have demonstrated that supra-
molecular hydrogels minimize the denaturation of proteins.14 Thus,
supramolecular hydrogels can serve as an equivalent of the
aggregates of small molecules in cellular environments for the
binding of the protein targets. Most importantly, the diameters
of the molecular nanofibers are on the nanometer scale, which is
comparable to the sizes of protein complexes. Such finite sizes
minimize the area of hydrophobic patches, thus reducing protein
denaturation caused by nonspecific interactions. In addition, it is
easy to separate the hydrogel particles from aqueous solution by
centrifugation after they interact with the proteins.
Driven by noncovalent interactions, molecular self-assembly—that
is, the spontaneous association of molecules results in structurally
well-defined aggregates (e.g. in the form of nanofibers, nano-
particles, or micelles)—is ubiquitous in biology.1 For example,
the self-assembly of proteins generates aggregates that are crucial
for cellular functions (e.g. actins and tubulins to form F-actins and
microtubules as cytoskeletons)2 or associate with certain diseases
(e.g. fibrillar aggregates of aberrant proteins as hallmarks of
neurodegenerative diseases),3 the self-assembly of lipids forms the
cell membrane that compartmentalizes subcellular organelles and
controls the substance in and out of cells,2 and the development of
small molecule hydrogels (e.g. lanreotide autogel) for treating
acromegaly.4 These essential processes of cells have led to intensive
studies of the aggregates of proteins for elucidating cellular
functions and mechanisms of certain diseases,5 as well as
exploration of the aggregates of small molecules for under-
standing the origin of life.6
Because one of the essential features of molecular self-assembly is
to exhibit emergent properties that drastically differ from those of
the individual molecules,1 the above well-established cases of
molecular self-assembly, thus, also raise an intriguing question
about the biological functions of the aggregates of small molecules
other than lipids, a question, however, remains unanswered. Three
major factors likely contribute to the lack of study of the biological
functions of the aggregates of small molecules: first, most of the
studies of the aggregates of small molecules, except the research on
lipids, focused on crystalline or liquid crystalline aggregates7
formed via molecular self-assembly in non-aqueous media.8 Despite
generating considerable physicochemical principles in the context
of supramolecular chemistry,9 those studies offer little insight for
the aggregates in biological processes that, obviously, occur in a
complex aqueous phase. Second, except the case of lipids, there are
As shown in Scheme 1, this assay consists of the following major
steps: (i) incubation of the hydrogel with cell lysate; (ii) photo
activation for fixing the proteins in the hydrogels; (iii) separation of
the hydrogel and removal of the non-specific proteins; (iv) analysis
and identification of the proteins (by SDS-PAGE, gel staining,
tandem mass spectrometry, and Western blotting, etc.). Our results
show that the nanofibers of a hydrophobic small peptide derivative
(1) exhibit rather selective interactions with tubulins and several
other proteins in the lysate of HeLa cells, which indicates that this
supramolecular hydrogel based protein binding assay can offer a
unique starting point for discovering interactions between the
molecular nanofibers and the proteins.
a Department of Chemistry, Brandeis University, 415 South Street,
Waltham, MA 02453, USA. E-mail: bxu@brandeis.edu;
Fax: +1 78 1736 2516; Tel: +1 78 1736 5201
b Department of Biology, Brandeis University, 415 South Street,
Waltham, MA 02453, USA
w Electronic supplementary information (ESI) available: Rheological
test, hydrogel washing conditions, additional electrophoresis results,
and synthesis. See DOI: 10.1039/c2cc33631f
To achieve the design in Scheme 1, we incorporate photoleucine,
a diazirine based photo reactive analog of leucine, into the
c
8404 Chem. Commun., 2012, 48, 8404–8406
This journal is The Royal Society of Chemistry 2012

View Article Online
Scheme 1 Illustration of the hydrogel protein pull-down assay
coupled with electrophoresis and tandem mass spectrometry for the
identification of the cytosolic proteins that bind to molecular nano-
fibers. The expanded box shows the photo reaction between the hydro-
gelator and the proteins bound to the molecular nanofibers.
Fig. 1 (A) Optical images of the hydrogel of 1 (0.6 wt%, pH 7.4)
without UV irradiation (left) and with UV irradiation (right). Typical
TEM images of the nanofibers in (B) the hydrogel of 1 with UV
irradiation; (C) the hydrogel of 1 without UV irradiation; (D) the hydrogel
of 1 incubated with cell lysate under UV irradiation and then thoroughly
washed. White arrow heads indicate the proteins on the nanofibers.
hydrogelator backbone because photo affinity is a successful direct
method to identify the interaction between a specific protein and its
biological target.15 The noncovalent interactions between the
hydrogelators allow the dissociation of the supramolecular
hydrogels, thus easily releasing the proteins bound to the
molecular nanofibers. We chose 1, which contains a naphthalene–
diphenylalanine (NapFF) motif, because the NapFF motif is able
to enable the self-assembly of many other bioactive molecules.16
Starting with naphthalene acetic acid, we connect it with Phe, Phe,
Lys(Boc), photo-Leu, and Tyr(P) sequentially via the cycle of NHS
activation and ester–amide exchange reaction and obtain the
precursor of the hydrogelator 1 after the deprotection of the lysine
via the cleavage of the Boc group (Scheme S1, ESIw). After
dissolving the precursor in water at a concentration of 0.6 wt%
(pH 7.4, adjusted by 1 M NaOH), we add 10 U mLꢀ1 of alkaline
phosphatase (ALP) to the precursor solution and obtain a
transparent hydrogel of 1 (Fig. 1A, left panel) after two hours
of dephosphorylation. The enzyme-induced formation of
hydrogelators16 readily ensures formation of the hydrogels
under the conditions that are compatible with cell lysate.
The negative staining transmission electron microscopy (TEM)
image reveals that the hydrogel of 1 contains the network of
uniform nanofibers that have a diameter of 10 ꢁ 2 nm (Fig. 1C).
To confirm the photo reactivity of the hydrogelator of 1, we use
UV light (365 nm) to irradiate the hydrogel. After UV irradiation,
the hydrogel becomes slightly yellowish (Fig. 1A, right panel). TEM
reveals that this light yellow hydrogel, though containing denser
nanofiber networks, consists of the nanofibers with the diameters of
10 ꢁ 2 nm (Fig. 1B). A rheological test indicates that the storage
moduli (G0) and loss moduli (G00) increase modestly after the UV
irradiation (Fig. S1, ESIw). After using UV to irradiate the hydrogel
of 1 incubated with the cell lysate and thoroughly washing the
irradiated hydrogel by PBS buffer containing 0.6% of Triton and
250 mM of NaCl, we use TEM to examine the hydrogel, and find
that protein particles and nanofibers co-exist in the hydrogel
(Fig. 1D). These results suggest that (i) the hydrogelator of 1 is
photo reactive; (ii) the interaction between the hydrogelators is
largely noncovalent despite the UV irradiation; (iii) UV irradiation
is able to retain the proteins on the nanofibers of 1.
the molecular nanofibers, we use a series of concentrations of a
Triton surfactant (as indicated on the top of the gel) together
with 250 mM of NaCl solution for washing the hydrogels after
the photoreaction. The purpose of the photoreaction is to
retain the proteins that bind to the nanofibers in the hydrogels.
After washing, we release the proteins from the hydrogels with
the addition of laemmli buffer to a final concentration of 2ꢂ.17
After being boiled for 4 minutes and centrifuged at 14 000 rpm
for 3 minutes, the sample was loaded onto the SDS-PAGE gel
for analysis. As shown in Fig. 2 (see Fig. S2 (ESIw) for the full
scope of the SDS-PAGE gel), with the increase in the
concentrations of Triton, most of the proteins desorb from
the hydrogels. Silver stain reveals that the intensities of the
bands of the non-specifically-absorbed proteins decrease with
the increase in the Triton concentration (e.g. the band indicated
by (o)). However, as shown in lanes 2 and 3 of Fig. 2, the
intensities of two protein bands (within the molecular weight
range of 37 to 75 kDa, indicated by (*)) remain almost constant
when the concentrations of the Triton are 0.6% and 0.3%,
implying that these two bands correspond to the proteins
interacting specifically with the nanofibers of 1.
To identify the proteins that interact with the molecular nano-
fibers of 1, the protein bands with 37–75 kDa are analyzed by in-gel
Fig. 2 Silver staining shows that different washing conditions alter
the ratio of proteins binding on the supramolecular hydrogel. Under
the harshest washing conditions, the hydrogels afford two major bands
(highlighted by * in dashed line boxes) while other bands fade away,
indicating the specific binding between those proteins and hydrogels.
The ratio of those two bands (top/bottom) is 3 : 7 in lane 2.
We perform the hydrogel-based protein pull-down assay
outlined in Scheme 1 and confirm that the nanofibers of 1 are
able to interact selectively with proteins. To find the proper
conditions to remove the nonspecifically-absorbed proteins on
c
This journal is The Royal Society of Chemistry 2012
Chem. Commun., 2012, 48, 8404–8406 8405

View Article Online
ultimately contribute to establish the molecular foundation
for the functions and applications of aggregates (e.g. nano-
fibers) of small molecules in the cellular environment.
This work was partially supported by NIH (R01CA142746)
and HFSP (RGP0056/2008). We thank EM facilities of Bran-
deis University for TEM. M. J. C. L. acknowledges the HHMI
predoctoral fellowship.
Notes and references
1 G. M. Whitesides and B. Grzybowski, Science, 2002, 295, 2418–2421.
2 H. Lodish, A. Berk, C. A. Kaiser, M. Krieger, M. P. Scott,
A. Bretscher, H. Ploegh and P. Matsudaira, Molecular Cell Biology,
W. H. Freeman, New York, 6th edn, 2007.
3 H. W. Querfurth and F. M. LaFerla, N. Engl. J. Med., 2010, 362,
329–344.
4 P. Caron, A. Beckers, D. R. Cullen, M. I. Goth, B. Gutt,
P. Laurberg, A. M. Pico, M. Valimaki and W. Zgliczynski,
J. Clin. Endocrinol. Metab., 2002, 87, 99–104.
5 J. R. Glover and S. Lindquist, Cell (Cambridge, Mass.), 1998, 94,
73–82.
6 J. W. Szostak, Philos. Trans. R. Soc. London, Ser. B, 2011, 366,
2894–2901; S. Koga, D. S. Williams, A. W. Perriman and S. Mann,
Nat. Chem., 2011, 3, 720–724.
7 L. Schmidt-Mende, A. Fechtenkotter, K. Mullen, E. Moons,
R. H. Friend and J. D. MacKenzie, Science, 2001, 293, 1119.
8 G. M. Whitesides, J. P. Mathias and C. T. Seto, Science, 1991, 254,
1312–1319.
9 J. M. Lehn, Angew. Chem., Int. Ed., 1988, 27, 89–112.
10 Z. H. Wang, H. M. Wang, W. T. Zheng, J. Zhang, Q. Zhao,
S. F. Wang, Z. M. Yang and D. L. Kong, Chem. Commun., 2011,
47, 8901–8903; H. Kobayashi, A. Friggeri, K. Koumoto, M. Amaike,
S. Shinkai and D. N. Reinhoudt, Org. Lett., 2002, 4, 1423–1426;
L. A. Estroff, L. Addadi, S. Weiner and A. D. Hamilton, Org.
Biomol. Chem., 2004, 2, 137–141; L. A. Estroff and A. D. Hamilton,
Chem. Rev., 2004, 104, 1201–1217; S. Ray, A. K. Das and
A. Banerjee, Chem. Commun., 2006, 2816–2818; S. Matsumoto,
S. Yamaguchi, A. Wada, T. Matsui, M. Ikeda and I. Hamachi,
Chem. Commun., 2008, 1545–1547; L. X. Xiao, C. Liu, J. H. Zhu,
D. J. Pochan and X. Q. Jia, Soft Matter, 2010, 6, 5293–5297; H. Cui,
T. K. Hodgdon, E. W. Kaler, L. Abezgauz, D. Danino,
M. Lubovsky, Y. Talmon and D. J. Pochan, Soft Matter, 2007, 3,
945–955; J. W. Sadownik and R. V. Ulijn, Chem. Commun., 2010, 46,
3481–3483; R. J. Williams, A. M. Smith, R. Collins, N. Hodson,
A. K. Das and R. V. Ulijn, Nat. Nanotechnol., 2009, 4, 19–24;
Y. J. Wang, B. Desbat, S. Manet, C. Aime, T. Labrot and R. Oda,
J. Colloid Interface Sci., 2005, 283, 555–564; N. M. Sangeetha and
U. Maitra, Chem. Soc. Rev., 2005, 34, 821–836.
11 Z. Yang, G. Liang and B. Xu, Acc. Chem. Res., 2008, 41, 315–326.
12 A. M. Kloxin, A. M. Kasko, C. N. Salinas and K. S. Anseth,
Science, 2009, 324, 59–63.
13 J. B. Matson and S. I. Stupp, Chem. Commun., 2012, 48, 26–33.
14 S. Kiyonaka, K. Sada, I. Yoshimura, S. Shinkai, N. Kato and
I. Hamachi, Nat. Mater., 2004, 3, 58–64.
15 M. Suchanek, A. Radzikowska and C. Thiele, Nat. Methods, 2005,
2, 261–267.
16 Y. Zhang, Y. Kuang, Y. A. Gao and B. Xu, Langmuir, 2011, 27,
529–537.
Fig. 3 (A) Tandem MS analysis indicates the most abundant hydrogel-
bound cytosolic proteins by sequence coverage. Black: proven binding
partners; grey partners that bind to cytoskeleton protein/fibers; white
proteins that were identified with no known association with the verified
proteins. Validation of protein–nanofibers interactions: (B) cell lysate was
incubated with the hydrogel and thoroughly washed under the conditions
shown on the top and subjected to Western blotting with antibody to actin;
(C) cell lysate was incubated with the hydrogel and thoroughly washed
under the conditions of 250 mM of NaCl, 0.6% triton, w/o colchicine, and
subjected to Western blotting with antibodies to tubulin and GAPDH
(another abundant protein expressed inside mammalian cells).
digestion18 and tandem MS analysis.19 Fig. 3A shows the proteins
that have the sequence coverage higher than 20% from the tandem
MS analysis. The sequence coverage of the proteins suggests that
the molecular nanofibers interact with tubulins, actins, and several
other proteins, which agrees well with the molecular weights of the
proteins in the bands of 55 kDa and 42 kDa.2
Based on the proteomic analysis (Fig. 3A), we use Western
blot20 to confirm some of the protein hits. As shown in Fig. 3B,
the protein bands at 42 kDa clearly contain actins. This result
also explains the observation of SEP-T7 in the MS spectra
because SEP-T7 can co-precipitate with actins.21 As a frequently
used control in protein analysis, Western blot of GAPDH
protein22 in Fig. 3C validates that the washing step successfully
removes nonspecific bound proteins from the hydrogel of 1. As
shown in Fig. 3C, Western blot proves the existence of tubulins
in the band of 55 kDa. The addition of 6 mM of colchicine (which
depolymerizes microtubules) hardly changes the binding results,
suggesting that the nanofibers bind both tubulins and micro-
tubules. This result also agrees with the high sequence coverage
of SEP-T2 because SEP-T2 can co-localize with microtubules.23
While the nature of the interactions between the nanofibers and
the other hits in Fig. 3A remains to be elucidated, we speculate
that a chaperonin (CCT-2) may interact with the peptidic
nanofibers of 1 because it binds to damaged or unfolded proteins.
In summary, this work indicates that the aggregates of small
molecules can selectively interact with certain protein targets.
Although the increase in the binding specificity of the nanofibers
toward a particular protein target relies on the molecular structure
of the small molecules, the negative result of anti-GAPDH blot in
Fig. 3C indicates that the interaction between tubulins and the
nanofibers of 1 is unlikely nonspecific. Because it is easy to
incorporate other bioactive molecules (e.g. taxol) into hydro-
gelators via the C-terminal16 or the e-amino group of lysine
residue24 of molecules like 1, this approach also allows the
study of self-assembled small molecule–protein interaction.
Thus, this supramolecular hydrogel pull-down assay offers a
facile way for elucidating the correlations between the protein
targets and the aggregates of the small molecules, which
17 Cold Spring Harb. Protoc., 2006, DOI: 10.1101/pdb.rec10424.
18 J. Rosenfeld, J. Capdevielle, J. C. Guillemot and P. Ferrara, Anal.
Biochem., 1992, 203, 173–179.
19 R. Aebersold and D. R. Goodlett, Chem. Rev., 2001, 101, 269–295.
20 J. Renart, J. Reiser and G. R. Stark, Proc. Natl. Acad. Sci. U. S. A.,
1979, 76, 3116–3120.
21 M. Kinoshita, S. Kumar, A. Mizoguchi, C. Ide, A. Kinoshita, T.
Haraguchi, Y. Hiraoka and M. Noda, Genes Dev., 1997, 11, 1535–1547.
22 R. D. Barber, D. W. Harmer, R. A. Coleman and B. J. Clark,
Physiol. Genomics, 2005, 21, 389–395.
23 E. T. Spiliotis, M. Kinoshita and W. J. Nelson, Science, 2005, 307,
1781–1785.
24 Y. Gao, Y. Kuang, Z. F. Guo, Z. H. Guo, I. J. Krauss and B. Xu,
J. Am. Chem. Soc., 2009, 131, 13576–13577.
c
8406 Chem. Commun., 2012, 48, 8404–8406
This journal is The Royal Society of Chemistry 2012