Nature has developed a vast array of post-translational
modifications that result in physical changes in protein
characteristics, functionality, and cellular location. An inge-
nious cellular mechanism for protein localization is prenyla-
tion: the covalent attachment of a hydrophobic prenyl group
to a protein to facilitate protein association with the plasma
membrane.[1] Analogous post-translational modifications that
induce protein–membrane anchorage include the covalent
attachment of glycolipid anchors (glycosylphosphatidylinosi-
tol, GPI)[2] and palmitoyl groups.[3] Through these processes,
otherwise soluble proteins are sequestered to cellular mem-
branes and ultimately lose their cellular motility.[4]
We postulated that the induced membrane anchorage of
proteins involved in cancer-promoting cell-signaling cascades
could hold significant therapeutic value. Thus, we wished to
explore the therapeutic potential of applying the principles of
protein anchorage to the development of a conceptually novel
drug modality. Our objective was to develop a scaffold that
could effectively restrict the motility of a cancer-promoting
protein within a cellular environment (Figure 1) and thereby
inhibit its function. Herein, we demonstrate the in vitro
application of this inhibition strategy and show the first
example of induced protein–membrane anchorage through
the use of a rationally designed protein–membrane anchor
(PMA).
To demonstrate this principle, we chose to target the
oncogenic signal transducer and activator of transcription 3
(STAT3) protein, a master regulator of the underlying events
in malignant transformation. Conceptually, our approach
represents a departure from traditional inhibitors of STAT3
cell-signaling pathways. Previous strategies have focused
primarily on the suppression of upstream kinases[5] and
STAT3–STAT3 protein–protein interactions.[6] To date,
these approaches have not yielded a clinically relevant
STAT3-targeting drug. STAT3 plays a key role in relaying
cytokine or growth-factor signaling to the nucleus, where it
binds to specific DNA-response elements in the promoters of
target genes and thereby induces cancer-promoting gene-
expression profiles. Thus, our goal was to develop an inhibitor
that could sequester STAT3, a 93 kDa protein, at the plasma
membrane and suppress nuclear translocation through PMA-
induced protein–membrane association.
Herein, we describe the design, synthesis, and application
of a novel PMA that targets STAT3 protein in liposome and
whole-cell systems. The prototype PMAwas composed of two
binding modules: a recognition motif to bind the protein
(STAT3) and an anchor to sequester the protein complex at
the membrane. Proof-of-concept PMA 1 (Figure 1) com-
prised a potent STAT3-recognition sequence GpYLPQTV-
NH2[7] covalently attached to a cholesterol membrane anchor
through the N terminus. The GpYLPQTV-NH2 peptide
sequence binds to the Src homology 2 (SH2) domain of
STAT3. It is the most potent STAT3 binder that has been
described and is an excellent handle for the coupling of our
lipid anchors.[6a] Owing to facile synthetic procedures and
potent membrane insertion, we elected to employ cholesterol
as our membrane anchor in preference to prenyl and GPI
lipids. Moreover, in support of this strategy, Simons and co-
workers have successfully used cholesterol to anchor drugs
that target membrane-embedded proteins to the plasma
membrane.[8] We attached the peptide to the cholesterol
unit in high yield through chloroformate coupling (see the
Supporting Information). Furthermore, to examine the role
played by the linking group, we prepared a PMA in which an
extended poly(ethylene glycol) linker (PEG) was used to link
the peptide and cholesterol moieties (PMA 2; Figure 1). As a
control compound, we synthesized a bivalent fluoresceinated
probe, 3, which incorporates a 5-aminofluorescein moiety
between the cholesterol unit and the peptide (Figure 1).
To evaluate whether the ditopic inhibitor 1 conserved its
STAT3-binding capability when conjugated to the cholesterol
steroid, we conducted control binding experiments with full-
length STAT3 protein. The binding affinity of 1 was measured
in a competitive fluorescence polarization (FP) assay popu-
larly used to determine the affinity of the STAT3 SH2 domain
for small molecules.[9] Encouragingly, we found that the
cholesterol conjugate retained good binding potency for the
STAT3 SH2 domain (Ki = 0.95 Æ 0.1 mm; GpYLPQTV-NH2:
Ki = 0.2 mm).
To assess the efficacy of our STAT3 PMAs, we developed
a series of in vitro fluorescence-based experiments to visual-
ize PMA-induced STAT3 protein localization in lipid model
systems. As part of these experiments, the cysteine thiol
groups in the protein were labeled with tetramethylrhod-
amine, and the resulting protein (TMR–STAT3) was charac-
terized by single-molecule spectroscopy (see the Supporting
Information).[10] To assess whether fluorescence labeling
compromised the phosphopeptide-binding function of the
STAT3 SH2 domain, we conducted control experiments on a
multiparameter confocal microscope.[11] Simultaneously
detected polarization and fluorescence correlation data con-
firmed that the binding affinity of TMR–STAT3 for the
inhibitor was similar to that of the unlabeled protein.[10] We
conducted in vitro liposome experiments to determine
whether PMAs could sequester a fluorescently labeled
93 kDa STAT3 protein at a lipid membrane. The experiments
were monitored with custom-built fluorescence microscopes
capable of the hyperspectral detection of single emitters.[11]
[*] M. Avadisian,[+] Dr. S. Fletcher,[+] B. Liu,[+] D. Badali,
Prof. Dr. C. C. Gradinaru, Prof. Dr. P. T. Gunning
Department of Chemical and Physical Sciences
University of Toronto Mississauga
Mississauga, ON L5L 1C6 (Canada)
Fax: (+1)905-828-5425
E-mail: claudiu.gradinaru@utoronto.ca
Dr. W. Zhao,[+] Dr. P. Yue, Prof. Dr. J. Turkson
Burnett School of Biomedical Sciences, College of Medicine
University of Central Florida (USA)
Dr. W. Xu, Dr. A. D. Schimmer
Ontario Cancer Institute/Princess Margaret Hospital (Canada)
[+] These authors contributed equally.
[**] Financial support was provided by the Leukemia and Lymphoma
Society of Canada (P.T.G., C.C.G., and J.T.), the NSERC (P.T.G.,
C.C.G.), the University of Toronto, a CIHR training award (B.L.), and
NIH grants CA106439 and CA128865 (P.T.G. and J.T.).
Supporting information for this article is available on the WWW
Angew. Chem. Int. Ed. 2011, 50, 6248 –6253
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim