Functional virus-based polymer-protein nanoparticles by atom transfer radical polymerization

Article abstract of DOI:10.1021/ja203286n

Viruses and virus-like particles (VLPs) are useful tools in biomedical research. Their defined structural attributes make them attractive platforms for engineered interactions over large molecular surface areas. In this report, we describe the use of VLPs as multivalent macroinitiators for atom transfer radical polymerization. The introduction of chemically reactive monomers during polymerization provides a robust platform for post-synthetic modification via the copper-catalyzed azide-alkyne cycloaddition reaction. These results provide the basis to construct nanoparticle delivery vehicles and imaging agents using protein-polymer conjugates.

Full text of DOI:10.1021/ja203286n

COMMUNICATION
pubs.acs.org/JACS
Functional Virus-Based PolymerꢀProtein Nanoparticles by Atom
Transfer Radical Polymerization
Jonathan K. Pokorski,† Kurt Breitenkamp,† Lars O. Liepold,‡ Shefah Qazi,‡ and M.G. Finn*,†
†Department of Chemistry and The Skaggs Institute for Chemical Biology, The Scripps Research Institute, La Jolla, California 92037,
United States
‡Department of Chemistry, Center for BioInspired Nanomaterials, Montana State University, Bozeman, Montana 59717, United States
S Supporting Information
b
techniques have previously been used to prepare well-defined
ABSTRACT: Viruses and virus-like particles (VLPs) are
polymerꢀprotein hybrid structures that maintain their biological
functions while gaining improved pharmacokinetic properties.9
useful tools in biomedical research. Their defined structural
attributes make them attractive platforms for engineered
interactions over large molecular surface areas. In this report,
we describe the use of VLPs as multivalent macroinitiators
for atom transfer radical polymerization. The introduction
of chemically reactive monomers during polymerization
provides a robust platform for post-synthetic modification
via the copper-catalyzed azideꢀalkyne cycloaddition reac-
tion. These results provide the basis to construct nanopar-
ticle delivery vehicles and imaging agents using proteinꢀ
polymer conjugates.
We expected ATRP to provide polymers with narrow molecular
weight distributions, yielding structurally uniform hybrid proteinꢀ
polymer nanoparticles from the monodisperse viral platform.
ATRP initiators were installed on the surface of Q β using a
two-step “click” protocol (Figure 1), which would allow other
functional units to be installed at the same time, if desired. First,
N-hydroxysuccinimide 3 was used to attach an azide group to
∼180 ( 30 of the 720 surface-accessible amine groups of the
particle (a value that is controlled by the selection of reaction
conditions for this step and determined experimentally by mass
spectrometry analysis of the denatured protein after mod-
ification). The resulting azido particle 3 was then functionalized
with a triglyme-based ATRP initiator (4) by copper-catalyzed
azideꢀalkyne cycloaddition (CuAAC) to form Qβ macroinitia-
tor 5.10 Polymers were grown from the multivalent initiator using
a 2,20-bipyridine/CuBr/CuBr2 catalyst system that was previously
shown to yield high-molecular-weight polymers grafted from
protein-based initiators in water.11 A degassed, aqueous copper
solution (0.1 mM final concentration) was added to a solution of
particle 3 and the commercially available monomer 6 or 7
(final concentrations, 1 mg/mL (0.4 μM) in VLP and 72 mM
in monomer). After overnight room-temperature reactions un-
der nitrogen, the polymerizations were terminated by exposure
to air and addition of EDTA. The resulting proteinꢀpolymer
conjugates were purified from the small-molecule reactants by
sucrose gradient centrifugation and ultrapelleting.
Size exclusion chromatography (SEC) and dynamic light
scattering (DLS) were used to determine changes in particle
size following ATRP. The former (Figure 2A) showed a shift in
elution volume from ∼12 min for unmodified Qβ particles to
9.5 min for the significantly larger poly(OEGMA)-coated pro-
duct. A similar shift was seen in electrophoretic mobility when the
particles were analyzed on a native agarose gel, where polymer-
coated particles remained near the baseline, a significant retarda-
tion compared to particle 5 (Figure 2D). DLS measurements
(Figure 2E,F) confirmed the increase in particle hydrodynamic
radius from 14 nm (Q β wild-type) to ∼24 nm for the polymer-
coated particles, with a narrow size distribution (polydispersity =
18%). The virus capsids, when visualized by TEM, were shown to
remain intact following ATRP (Figure 2B,C).
ynthetic polymers are chemically diverse in terms of their size
S
and composition and have long been used for the display of
multiple copies of functional units. In contrast to their synthetic
counterparts, biopolymers such as viruses and virus-like particles
(VLPs) exhibit unique qualities of monodispersity and chemical
regularity, allowing for the precise, periodic chemical functiona-
lization of capsid structures.1 VLPs can be both genetically2 and
chemically1b modified to alter or introduce functionality and
tailor biological functions. The combination of these two types of
multivalent structures may prove advantageous in situations
requiring organic nanoparticles with defined structural attributes.
Synthetic polymer nanoparticles prepared by emulsion,
nanoprecipitation, self-assembly, or layer-by-layer techniques have
been widely studied as delivery vehicles.3 Their potential advan-
tages include high drug-loading capacities, ability to improve drug
solubility, and the ready introduction of ligands for targeted
delivery.4 However, the production of polymeric nanoparticles
with precise structural homogeneity remains a challenge to the
field.5 The use of biomolecular platforms is an attractive approach
toward this end, and many examples exist of the modification of
viral and non-viral protein structures with synthetic polymers
using “grafting to” approaches.6 In contrast, there are few examples
of the use of protein nanoparticles as multivalent macroinitiators
for polymerization,7 and none using viruses.8
We describe here the polymerization of oligo(ethylene
glycol)-methacrylate (6, OEGMA) and its azido-functionalized
analogue (7, OEGMA-N3) directly from the outer surface of the
bacteriophage Q β VLP by atom transfer radical polymerization
(ATRP). ATRP and other controlled radical polymerization
Received: April 10, 2011
Published: May 31, 2011
r
2011 American Chemical Society
9242
dx.doi.org/10.1021/ja203286n J. Am. Chem. Soc. 2011, 133, 9242–9245
|

Journal of the American Chemical Society
COMMUNICATION
Figure 3. End labeling of Qβ-poly(OEGMA) (8). For the structure of
11, see Figure 4.
Figure 1. Preparation of the Qβ VLP macroinitiator and polymeriza-
tion from its surface.
Figure 4. Conjugation to polymer side chains. (A) CuAAC reaction to
append functionality to 9. (B) UV illumination of sucrose gradient
following CuAAC between 9 and 11 (upper band, unreacted 11; lower
band, particle 14). (C) Conjugation of GdꢀDOTA complex 12 to
azide-functionalized capsid 3.
Figure 2. Characterization of Qβꢀpolymer particles. (A) Size exclu-
sion chromatographs. (B,C) Transmission electron micrographs of
poly(OEGMA)- and poly(OEGMA-N3)-coated particles (8 and 9,
respectively); scale bar = 200 nm. (D) Non-denaturing agarose gel
stained with ethidium bromide: lane 1, Qβ macroinitiator (5); lane 2,
Qβꢀpoly(OEGMA) (8); lane 3, Qβꢀpoly(OEGMA-N3) (9). (E,F)
DLS histograms of 8 and 9, respectively.
expect that the method of post-polymerization functionalization
and bioconjugation described here will prove especially useful in
attaching functional biomolecules such as cell targeting groups,
cell penetrating peptides, or diagnostic agents.
The azido-functionalized oligo(ethylene glycol)-methacrylate,
OEGMA-N3 (7), also served as a competent monomer, allowing
for a greater degree of post-polymerization functionalization while
retaining PEG-like biocompatibility.13 The polymerization of 7
proceeded in the same manner as 6 (Figure 1), yielding hybrid
Q βꢀpoly(OEGMA-N3) particle 9 with properties very similar
to those of the poly(OEGMA)-coated 8: ∼10 nm increase in
radius, narrow size distribution, and dramatically different elec-
trophoretic mobility (Figure 2A,C,D,F).
To demonstrate the reactivity of poly(OEGMA-N3) grafts, 9
was reacted with alkyne-substituted AlexaFluor488 dye 11 under
CuAAC conditions (Figure 4A). Sucrose gradient centrifugation
revealed an intensely colored green band, indicating a successful
“click” reaction (Figure 4B). The potential sensitivity of the
molar absorptivity of these dyes to their molecular environment
made quantification of the coupling efficiency at high dye loadings
impossible by absorbance measurements. The same reaction was
therefore performed with gadolinium complex11, and the coupling
result assayed by quantitation of Gd by inductively coupled
plasma optical emission spectroscopy (ICP-OES).14 Loading
values of 500ꢀ650 Gd complexes per particle were routinely
found for reactions with different samples of 9.
ATRP provides a tertiary bromide at the terminus of
each growing polymer chain, which is amenable to further
transformation.12 As shown in Figure 3, we reacted particle 8,
which does not contain side-chain azide functionality, with a large
molar excess of sodium azide in a water:DMSO mixture to
substitute azide for bromide at the particle periphery. The
resulting intermediate was then reacted with AlexaFluor488-
alkyne under standard CuAAC bioconjugation conditions. Fol-
lowing sucrose gradient purification, the fluorescent particles
were found to bear an average of ∼20 dye molecules per particle
by comparing Alexa UV absorbance (495 nm) versus total
protein concentration (as measured by Bradford assay). No
observable dye labeling occurred when 8 was subjected directly
to CuAAC prior to sodium azide treatment, suggesting that no
accessible azides remained following CuAAC to particle 3. The
labeling of only 20 chains on a particle that started with 180
potential initiation sites suggests that either polymer initiation is
strongly inhibited by previously initiated chains or that azide
substitution and click cycloaddition reactions are inhibited by the
presence of adjacent chains attached to the particle surface. Given
the high efficiency of both azide nucleophilic attack and CuAAC
coupling, we suspect that the former explanation is more likely.
This issue will be studied further in upcoming experiments. We
Nanoparticles bearing Gd complexes have attracted substan-
tial interest as magnetic resonance imaging agents due to benefits
anticipated for their large size (slowing rotational relaxation
9243
dx.doi.org/10.1021/ja203286n |J. Am. Chem. Soc. 2011, 133, 9242–9245

Journal of the American Chemical Society
COMMUNICATION
Table 1. T1 Relaxivities (mMꢀ1 sꢀ1) for Derivatized Virus-
like Particles
relaxivity per Gd (per VLP)
sample
Gd/VLP
60 MHz
90 MHz
15
17
610
350
11.63 (7092)
10.7 (3750)
10.12 (6174)
11.64 (4075)
Figure 5. (A) In vitro release profile of doxorubicin from particle 16 at
pH 7.4 and 5.5 (MES = 2(N-morpholino)ethanesulfonic acid buffer;
PBS = phosphate-buffered saline). (B) MTT assay for cell viability after
treatment with the indicated agents; see text.
rates) and high Gd loading capacities. Several variations using VLPs
have been reported.15 We had earlier described the direct attach-
ment of Gd(DOTA)ꢀalkyne complex 12 to azide-derivatized
particle 3 and the magnetic resonance behavior of the resulting
conjugate 17.14 Examination of the QβꢀpolymerꢀGd particle 15
found a relaxivity on a per Gd basis similar to that of 17 at two
different Larmor frequencies (Table 1; approximately double the
value of the Gd complex alone), suggesting that the polymer
backbone does not impart additional conformational rigidity to the
complexes or restrict access to water. The per particle relaxivity of
15 (7092 mMꢀ1 sꢀ1), however, compares favorably to examples
reported in the literature involving the covalent attachment of
commercially available Gd reagents to virus particles.15bꢀd
The multiple attachment points offered by the azide groups of
polymer-decorated particle 9 may also be used to tether a re-
leasable drug cargo. It has been reported that nanoparticles of
∼25 nm radius are optimal for internalization via endocytotic
pathways.16 The relatively acidic environments of endosomal
and lysosomal compartments (pH 4.5ꢀ5.5) is most often used as
the trigger for drug release, with hydrazones being a popular
motif.17 Accordingly, we prepared a “clickable” doxorubicin
hydrazone13 for conjugation to the Qβꢀpolyazide 9 (Figure4A).
Because 13 has poorer aqueous solubility than the Gd complex,
fewer doxorubicin (Dox) molecules were attached, an average of
150 per particle (∼3 wt %), determined by measuring Dox
concentrations spectrophotometrically (λabs = 490 nm) and
comparing these to protein concentrations obtained by Bradford
assay.18 The resulting particles 16 remained intact, with a narrow
size distribution (by DLS, radius = 22.7 nm, 14% polydispersity).
This stands in contrast to the conjugation of 13 directly to
particle 3, which led to degradation of the capsid structure and
precipitation of the protein, as occasionally happens with hydro-
phobic or polyaromatic molecules.
The pH sensitivity of the Dox-conjugated particle 16 was
assayed by fluorescence spectroscopy. The fluorescence of the
Dox chromophores on 16 is known to be quenched at high local
concentrations,19 allowing us to monitor the release of the drug
(λex = 480 nm; λem = 598 nm) from the conjugate into solution at
pH 7.4 and 5.5.20 The maximum fluorescence was reached after
∼2 h at pH 5.5, while no fluorescence increase was observed after
12 h for the particles incubated at pH 7.4 (Figure 5A), consistent
with the expected properties of the hydrazone linkage.
The pH-sensitive, Dox-conjugated particle 16 was also found
to be cytotoxic to HeLa cells, a cervical cancer cell line that is
sensitive to treatment with Dox.21 Cells were incubated with varying
concentrations of 16 or appropriate controls for 8 h in pH 7
buffer (in which no cleavage of Dox from the particle occurs) and
were assayed for cell viability the following day using the MTT
assay. Polymer conjugate 16 exhibited a cytotoxicity profile similar
to that of free Dox (in terms of overall Dox concentrations)
under the same assay conditions (Figure 5B), indicating that the
particles are internalized and release their payload to effect cell
death. The cells showed no significant signs of toxicity when
exposed to unloaded particles (9) at the concentration range
used with 16 (Supporting Information).
In summary, we have shown that VLPs derived from the
bacteriophage Qβ can serve as a platform for controlled radical
polymerization to produce polymer-coated protein nanoparti-
cles. The size and surface properties of Qβ can be significantly
altered through ATRP while still retaining the low polydisper-
sities associated with VLPs. Furthermore, the polymer-coated
particles are accessible to bioconjugation at both the chain termini
and suitably derivatized polymer side chains. The platform allows
for both conjugation of small-molecule imaging agents and
chemotherapeutics. Future studies will focus on varying the
nature and properties of the polymers grown from capsid
surfaces and on further exploration of this platform for develop-
ment of targeted drug delivery and imaging agents.
’ ASSOCIATED CONTENT
S
Supporting Information. Detailed procedures for all
b
chemical transformations and cell-based assays. This material is
available free of charge via the Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION
Corresponding Author
mgfinn@scripps.edu
’ ACKNOWLEDGMENT
This work was supported by the National Institutes of Health
(CA112075, RR021886, R21EB005364) and by a Pathway to
Independence Award (K99EB011530) to J.K.P. We thank Prof.
Trevor Douglas for helpful discussions and Dr. Burkhardt Laufer
for the synthesis of Gd complex 12.
’ REFERENCES
(1) (a) Steinmetz, N. F. Nanomedicine: Nanotechnol., Biol, Med.
2010, 6, 634–641. (b) Strable, E.; Finn, M. G. Curr. Top. Microbiol.
Immunol. 2009, 327, 1–21.
(2) Brown, S.; Fiedler, J.; Finn, M. G. Biochemistry 2009, 48, 11155–
11157.
(3) Mora-Huertas, C. E.; Fessi, H.; Elaissari, A. Int. J. Pharm. 2010,
385, 113–142.
(4) Petros, R. A.; Desimone, J. M. Nat. Rev. Drug Discovery 2010,
9, 615–627.
(5) Good control over size has been achieved in many systems,
including polymer nanoparticles: (a) Duan, H.; Kuang, M.; Zhang, G.;
Wang, D.; Kurth, D. G.; M€ohwald, H. Langmuir 2005, 21, 11495–11499.
(b) Yanagishita, T.; Fujimura, R.; Nishio, K.; Masuda, H. Langmuir
9244
dx.doi.org/10.1021/ja203286n |J. Am. Chem. Soc. 2011, 133, 9242–9245

Journal of the American Chemical Society
COMMUNICATION
2009, 26, 1516–1519. (c) An, S. Y.; Bui, M.-P. N.; Nam, Y. J.; Han, K. N.;
Li, C. A.; Choo, J.; Lee, E. K.; Katoh, S.; Kumada, Y.; Seong, G. H.
J. Colloid Interface Sci. 2009, 331, 98–103.
(6) (a) Kovacs, E. W.; Hooker, J. M.; Romanini, D. W.; Holder,
P. G.; Berry, K. E.; Francis, M. B. Bioconjugate Chem. 2007,
18, 1140–1147. (b) Brunel, F. M.; Lewis, J. D.; Destito, G.; Steinmetz,
N. F.; Manchester, M.; Stuhlmann, H.; Dawson, P. E. Nano Lett. 2010,
10, 1093–1097. (c) Bruckman, M. A.; Kaur, G.; Lee, L. A.; Xie, F.;
Sepulveda, J.; Breitenkamp, R.; Zhang, X.; Joralemon, M.; Russell, T. P.;
Emrick, T.; Wang, Q. ChemBioChem 2008, 9, 519–523. (d) Holder,
P. G.; Finley, D. T.; Stephanopoulos, N.; Walton, R.; Clark, D. S.;
Francis, M. B. Langmuir 2010, 26, 17383–17388.
(7) (a) Hu, Y. X.; Samanta, D.; Parelkar, S. S.; Hong, S. W.; Wang,
Q. A.; Russell, T. P.; Emrick, T. Adv. Funct. Mater. 2010, 20, 3603–3612.
(b) Zeng, Q.; Li, T.; Cash, B.; Li, S.; Xie, F.; Wang, Q. Chem. Commun.
2007, 1453–1455.(c) Li, H.; Li, M.; Yu, X.; Bapat, A. P.; Sumerlin, B. S.
Polym. Chem. 2011, DOI 10.1039/c1py00031d.
(8) Many examples exist showing polymerization from the surfaces of
polymer and metal nanoparticles. For examples, see: (a) Barbey, R.; Lavanant,
L.;Paripovic, D.;Schuwer, N.;Sugnaux, C.;Tugulu, S.;Klok, H.-A.Chem. Rev.
2009, 109, 5437–5527. (b) Zou, H.; Wu, S.; Shen, J. Chem. Rev. 2008,
108, 3893–3957. For key examples, see:(c) Ali, A. M. I.; Mayes, A. G.
Macromolecules 2010, 43,837–844. (d) Skaff, H.;Emrick, T.Angew. Chem. Int.
Ed. 2004, 43, 5383–5386. (e) Perruchot, C.; Khan, M. A.; Kamitsi, A.; Armes,
S. P.; Von Werne, T.; Patten, T. E. Langmuir 2001, 17, 4479–4481.
(9) (a) Heredia, K. L.; Bontempo, D.; Ly, T.; Byers, J. T.; Halstenberg,
S.; Maynard, H. D. J. Am. Chem. Soc. 2005, 127, 16955–16960. (b)
Bontempo, D.; Maynard, H. D. J. Am. Chem. Soc. 2005, 127, 6508–6509.
(c) De, P.; Li, M.; Gondi, S. R.; Sumerlin, B. S. J. Am. Chem. Soc. 2008,
130, 11288–11289. (d) Lele, B. S.; Murata, H.; Matyjaszewski, K.; Russell,
A. J. Biomacromolecules 2005, 6, 3380–3387.
(10) (a) Hong, V.; Presolski, S. I.; Ma, C.; Finn, M. G. Angew. Chem.
Int. Ed. 2009, 48, 9879–9883.(b) Control experiments showed that no
degradation of or insertion into the tertiary bromide initiation site occurs
under CuAAC reaction conditions.
(11) Peeler, J. C.; Woodman, B. F.; Averick, S.; Miyake-Stoner, S. J.;
Stokes, A. L.; Hess, K. R.; Matyjaszewski, K.; Mehl, R. A. J. Am. Chem.
Soc. 2010, 132, 13575–13577.
(12) (a) Golas, P.; Tsarevsky, N.; Sumerlin, B.; Walker, L.;
Matyjaszewski, K. Aust. J. Chem. 2007, 60, 400–404. (b) Tsarevsky, N. V.;
Sumerlin, B. S.; Matyjaszewski, K. Macromolecules 2005, 38, 3558–3561.
(13) (a) Gao, W.; Liu, W.; Christensen, T.; Zalutsky, M.; Chilkoti, A.
Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 16432–16437. (b) Gao, W.; Liu,
W.; Mackay, J. A.; Zalutsky, M. R.; Toone, E. J.; Chilkoti, A. Proc. Natl.
Acad. Sci. U.S.A. 2009, 106, 15231–15236.
(14) Prasuhn, D. E.; Yeh, R. M.; Obenaus, A.; Manchester, M.; Finn,
M. G. Chem. Commun. 2007, 1269–1271.
(15) (a) Liepold, L. O.; Abedin, M. J.; Buckhouse, E. D.; Frank, J.;
Young, M. J.; Douglas, T. Nano Lett. 2009, 9, 4520–4526. (b) Liepold,
L.; Anderson, S.; Willits, D.; Oltrogge, L.; Frank, J.; Douglas, T.; Young,
M. Magn. Reson. Med. 2007, 58, 871–879. (c) Hooker, J. M.; Datta, A.;
Botta, M.; Raymond, K. N.; Francis, M. B. Nano Lett. 2007,
7, 2207–2210. (d) Anderson, E. A.; Isaacman, S.; Peabody, D. S.; Wang,
E. Y.; Canary, J. W.; Kirshenbaum, K. Nano Lett. 2006, 6, 1160–1164.
(16) Zhang, S.; Li, J.; Lykotrafitis, G.; Bao, G.; Suresh, S. Adv. Mater.
2009, 21, 419–424.
(17) (a) De Jesus, O. L. P.; Ihre, H. R.; Gagne, L.; Frechet, J. M. J.;
Szoka, F. C. Bioconjugate Chem. 2002, 13, 453–461. (b) Bae, Y.; Fukushima,
S.; Harada, A.; Kataoka, K. Angew. Chem. Int. Ed. 2003, 42, 4640–4643. (c)
Ulbrich, K.; Subr, V. Adv. Drug Delivery Rev. 2004, 56, 1023–1050.
(18) Missirlis, D.; Kawamura, R.; Tirelli, N.; Hubbell, J. Eur. J. Pharm.
Sci. 2006, 29, 120–129.
(19) Mohan, P.; Rapoport, N. Mol. Pharmaceutics 2010, 7, 1959–1973.
(20) (a) Ta, T.; Convertine, A. J.; Reyes, C. R.; Stayton, P. S.; Porter,
T. M. Biomacromolecules 2010, 11, 1915–1920. (b) Karukstis, K. K.;
Thompson, E. H. Z.; Whiles, J. A.; Rosenfeld, R. J. Biophys. Chem. 1998,
73, 249–263.
(21) Eichholtz-Wirth, H. Br. J. Cancer 1980, 41, 886–891.
9245
dx.doi.org/10.1021/ja203286n |J. Am. Chem. Soc. 2011, 133, 9242–9245