Design of Environmentally Responsive Fluorescent Polymer Probes for Cellular Imaging

Article abstract of DOI:10.1021/acs.biomac.5b00591

We report the development of environmentally responsive fluorescent polymers. The reversible temperature-induced phase transition of copolymers composed of N-isopropylacrylamide and a fluorescent monomer based on the fluorescein (FL), coumarin (CO), rhodamine (RH), or dansyl (DA) skeleton was used as a molecular switch to control the fluorescence intensity. The poly(N-isopropylacrylamide) (PNIPAAm) chain showed an expanded coil conformation below the lower critical solution temperature (LCST) due to hydration, but it changed to a globular form above the LCST due to dehydration. Through the combination of a polarity-sensitive fluorophore with PNIPAAm, the synthetic fluorescent polymer displayed a response to external temperature, with the fluorescence strength dramatically changing close to the LCST. Additionally, the P(NIPAAm-co-FL) and P(NIPAAm-co-CO) polymers, containing fluorescein and coumarin groups, respectively, exhibited pH responsiveness. The environmental responsiveness of the reported polymers is derived directly from the PNIPAAm and fluorophore structures, thus allowing for the cellular uptake of the fluorescence copolymer by RAW264.7 cells to be temperature-controlled. Cellular uptake was suppressed below the LCST but enhanced above the LCST. Furthermore, the cellular uptake of both P(NIPAAm-co-CO) and P(NIPAAm-co-RH) conjugated with a fusogenic lipid, namely, L-α-phosphatidylethanolamine, dioleoyl (DOPE), was enhanced. Such lipid-conjugated fluorescence probes are expected to be useful as physiological indicators for intracellular imaging. (Chemical Equation Presented).

Full text of DOI:10.1021/acs.biomac.5b00591

Article
pubs.acs.org/Biomac
Design of Environmentally Responsive Fluorescent Polymer Probes
for Cellular Imaging
Arisa Yamada, Yuki Hiruta, Jian Wang, Eri Ayano, and Hideko Kanazawa*
Faculty of Pharmacy, Keio University, 1-5-30, Shibakoen, Minato-ku, Tokyo 105-8512, Japan
S
* Supporting Information
ABSTRACT: We report the development of environmentally
responsive fluorescent polymers. The reversible temperature-
induced phase transition of copolymers composed of N-
isopropylacrylamide and a fluorescent monomer based on the
fluorescein (FL), coumarin (CO), rhodamine (RH), or dansyl
(DA) skeleton was used as a molecular switch to control the
fluorescence intensity. The poly(N-isopropylacrylamide)
(PNIPAAm) chain showed an expanded coil conformation
below the lower critical solution temperature (LCST) due to
hydration, but it changed to a globular form above the LCST
due to dehydration. Through the combination of a polarity-
sensitive fluorophore with PNIPAAm, the synthetic fluores-
cent polymer displayed a response to external temperature,
with the fluorescence strength dramatically changing close to the LCST. Additionally, the P(NIPAAm-co-FL) and P(NIPAAm-
co-CO) polymers, containing fluorescein and coumarin groups, respectively, exhibited pH responsiveness. The environmental
responsiveness of the reported polymers is derived directly from the PNIPAAm and fluorophore structures, thus allowing for the
cellular uptake of the fluorescence copolymer by RAW264.7 cells to be temperature-controlled. Cellular uptake was suppressed
below the LCST but enhanced above the LCST. Furthermore, the cellular uptake of both P(NIPAAm-co-CO) and P(NIPAAm-
co-RH) conjugated with a fusogenic lipid, namely, L-α-phosphatidylethanolamine, dioleoyl (DOPE), was enhanced. Such lipid-
conjugated fluorescence probes are expected to be useful as physiological indicators for intracellular imaging.
based polymer have been studied extensively.7−9 Although a
range of fluorescent polymers has been developed, the majority
INTRODUCTION
■
Environmentally responsive polymers are well-known as
allow only simple monitoring of the solution temperature in
valuable materials in biomaterial sciences and biomedical
technologies.1 Such polymers change their structures and
terms of emission intensity.
Recently, we developed a dual temperature/pH-responsive
fluorescent polymer for application in cellular imaging.10 The
temperature and pH of all living cells are known to change
during cellular events such as enzyme reactions, gene
expression, and metabolism.11 From a clinical viewpoint,
pathological cells are warmer than normal cells due to their
enhanced metabolic activity.12 Abnormal intracellular pH values
are therefore associated with inappropriate cell function and
growth and are observed in a number of common diseases such
as cancer13 and Alzheimer’s disease.14 Monitoring intracellular
temperature and pH, therefore, can provide critical information
for studying physiological and pathological processes within
living cells. We previously reported the synthesis of poly(N-
isopropylacrylamide) (PNIPAAm) bearing a fluorescent
molecule at the terminal end and confirmed its temperature-
and pH-dependent cellular uptake.15,16
physical properties rapidly and reversibly in response to subtle
external stimuli such as temperature, pH, and light. Within this
group of polymers, poly(N-isopropylacrylamide) (PNIPAAm)
has been found to exhibit thermally reversible solubility changes
in response to temperature changes across its lower critical
solution temperature (LCST) at 32 °C.2 In aqueous solution,
PNIPAAm shows an expanded coil conformation below the
LCST due to strong hydration, but it changes to a globular
form above the LCST due to sudden dehydration. This unique
property has allowed PNIPAAm to be widely applied in drug
delivery systems,3 cell culture substrates,4 and separation
systems.5
Recently, fluorescent polymers composed of environmentally
responsive polymers and fluorescent molecules have received a
great deal of attention. It is well-known that the wavelength and
strength of a fluorescent molecule are sensitive to changes in
the adjacent molecular environment, including density, temper-
ature, and pH. Annaka et al. reported the use of a PNIPAAm-
based fluorescent polymer to study the swelling behavior of
grafted PNIPAAm hydrogels by fluorescence depolarization.6
In addition, fluorescent thermometer beads on a PNIPAAm-
Received: May 1, 2015
Revised: June 17, 2015
© XXXX American Chemical Society
A
DOI: 10.1021/acs.biomac.5b00591
Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules
Article
of their aqueous solution (0.5% w/v). The optical transmittance of
each polymer solution was measured at 500 nm over a range of
temperatures using a UV−vis spectrophotometer (V-630, JASCO,
Tokyo, Japan). The temperature was controlled using an ETC-717
With the aim of implementation in cellular environmental
imaging, we herein report the preparation of four fluorescent
PNIPAAm-based copolymers and characterization of the effects
of pH and/or temperature on their fluorescent behavior.
controller (JASCO) and a PT-31 Peltier system (Kruss, Hamburg,
̈
Germany); the heating rate was 0.1 °C/min. The LCST was
determined at the temperature at which 50% optical transmittance
of the polymer solution was achieved. Fluorescence spectra were
measured using an FP-6300 spectrofluorometer (JASCO), and the
temperature was controlled as described above. A quartz cuvette with a
1 cm path length was used. The maximum excitation wavelengths and
measured emission wavelengths of the fluorescent polymers were as
follows (0.5% w/v aqueous solution): P(NIPAAm-co-DA) λex = 310
nm, λem = 500 nm; P(NIPAAm-co-FL) λex = 440 nm, λem = 515 nm;
P(NIPAAm-co-CO) λex = 350 nm, λem = 460 nm; and P(NIPAAm-co-
RH) λex = 540 nm, λem = 590 nm. The effects of pH and temperature
on the fluorescence intensity of P(NIPAAm-co-FL) and P(NIPAAm-
co-CO) were evaluated in Briton−Robinson universal buffers (pH 3−
11) between 25 and 35 °C.
Synthesis of the Lipid-Conjugated Polymers. The terminal
carboxyl groups on P(NIPAAm-co-CO) and P(NIPAAm-co-RH) were
esterified using NHS in the presence of DCC (molar ratio, 1:2.5:2.5)
in dichloromethane prior to conjugation with DOPE. The activated
esterified polymer (200 mg) was dissolved in anhydrous 1,4-dioxane
(10 mL) and reacted with DOPE (25 mg) for 2 days at 25 °C. After
evaporation of the reaction solution, the residue was dissolved in
MeOH and subjected to dialysis, using a dialysis membrane with a
3500 amu cutoff (Spectra/Por, Spectrum Laboratories, CA, USA) at 4
°C for 3 days. After this time, evaporation of the solvent gave the
desired product as a white ([P(NIPAAm-co-CO)-DOPE]) or pink−
red solid ([P(NIPAAm-co-RH)-DOPE]).
MATERIALS AND METHODS
■
Materials. N-Isopropylacrylamide (NIPAAm) was kindly provided
by KJ Chemicals Corporation (Tokyo, Japan) and was purified by
recrystallization from n-hexane and dried at 25 °C in vacuo.
Fluorescein o-acrylate was purchased from Sigma-Aldrich (St. Louis,
MO, USA). 7-(4-Trifluoromethyl)coumarin acrylamide, dansyl
chloride, N,N-dimethylethylenediamine, acryloyl chloride, and triethyl-
amine were purchased from Tokyo Kasei Industry (Tokyo, Japan).
Methacryloxyethyl thiocarbamoyl rhodamine B was purchased from
Funakoshi (Tokyo, Japan). 2,2′-Azobis(isobutyronitrile) (AIBN), 3-
mercaptopropionic acid (MPA), and L-α-phosphatidylethanolamine,
dioleoyl (DOPE) were purchased from Wako (Osaka, Japan). N-
Hydroxysuccinimide (NHS) and N,N′-dicyclohexylcarbodiimide
(DCC) were purchased from Kanto Chemical (Tokyo, Japan). All
other reagents and solvents were of analytical grade.
Synthesis of N-[2-[[[5-(N,N-Dimethylamino)-1-
naphthalenyl]sulfonyl]amino]ethyl]-2-propenamide. N-[2-
[[[5-(N,N-Dimethylamino)-1-naphthalenyl]sulfonyl]amino]ethyl]-2-
propenamide (DA) was prepared according to literature methods.17 A
mixture of dansyl chloride (200 mg, 0.74 mmol) and N,N-
dimethylethylenediamine (445 mg, 7.42 mmol) in dichloromethane
was stirred at 0 °C for 1 h to give the DA precursor, N-(2-
aminoethyl)-5-(dimethylamino)naphthalene-1-sulfonamide. After pu-
rification by column chromatography (silica gel), N-(2-aminoethyl)-5-
(dimethylamino)naphthalene-1-sulfonamide (120 mg, 0.40 mmol) was
reacted with acryloyl chloride (45 mg, 0.49 mmol) in the presence of
triethylamine (50 mg, 0.49 mmol) in THF at 0 °C for 24 h. The crude
material was purified by column chromatography to give DA as a light
green solid (82 mg, 0.23 mmol). δ 1H NMR (600 MHz, CDCl3): 8.56
(d, 1H, J = 8.5 Hz), 8.24 (m, 2H), 7.55 (m, 2H), 7.19 (d, 1H, J = 7.3
Hz), 6.20(d, 1H, J = 16.9 Hz), 5.91 (m, 2H), 5.61 (d, 1H, J = 10.2
Hz), 5.31 (s, 1H), 3.37 (m, 2H), 3.07 (m, 2H), 2.90 (s, 6H).
Synthesis of Poly(N-isopropylacrylamide-co-DA). Poly(N-
isopropylacrylamide-co-DA) [P(NIPAAm-co-DA)] was prepared
using radical polymerization, as shown in Scheme 1. NIPAAm (10.0
g, 88.4 mmol) and DA (30.5 mg, 0.09 mmol) were dissolved in N,N-
dimethylformamide (20 mL). AIBN (58 mg, 0.35 mmol) and MPA
(263 mg, 2.48 mmol), which act as the radical initiator and the chain
transfer agent, respectively, were added to the solution. The reaction
mixture was degassed by subjecting to freeze−thaw cycles and then
heated at 70 °C for 5 h. The reaction solution was then poured into
diethyl ether (1 L) to precipitate the polymers. The crude product was
further purified by repeated precipitation using diethyl ether (1 L)
from a solution of acetone (20 mL), filtered, and dried to give the
desired polymer as a red solid (7.0 g). The molecular weight of the
polymer was determined by GPC analysis.
Synthesis of Other Fluorescence Polymers. Poly(N-isopropyl-
acrylamide-co-fluorescein o-acrylate) [P(NIPAAm-co-FL)], poly[N-
isopropylacryamide-co-7-(4-trifluoromethyl)coumarin acrylamide] [P-
(NIPAAm-co-CO)], and poly[N-isopropylacrylamide-co-methacryloxy-
ethyl thiocarbamoyl rhodamine B] [P(NIPAAm-co-RH)] were
prepared from NIPAAm and either fluorescein o-acrylate, 7-(4-
trifluoromethyl)coumarin acrylamide, or methacryloxyethyl thiocarba-
moyl rhodamine B, respectively, according to the above procedure.
Analytical Techniques. 1H NMR spectra were acquired on a
JNM-EP600 spectrometer (600 MHz, JEOL, Tokyo, Japan) using
tetramethylsilane as the internal standard. GPC was conducted on a
TOSOH GPC-8020 system equipped with a differential refractive
index detector, TSK guard column, and two TSK GEL α-M columns.
The mobile phase was composed of 10 mM LiCl in DMF (at 40 °C,
flow rate = 1.0 mL/min). Calibration was performed using near-
monodisperse poly(ethylene glycol) standards obtained from TOSOH
(Tokyo, Japan). The lower critical solution temperatures (LCSTs) of
the polymers were determined by measuring the optical transmittance
Cell Culture. RAW264.7 cells (RIKEN BRC Cell Bank) were
cultured as subconfluent monolayers in a 75 cm2 culture flask with a
vent cap using MEM, supplemented with 10% fetal bovine serum
(FBS), 50 units/mL penicillin, and 50 μg/mL streptomycin, at 37 °C
in a humidified incubator containing 5% CO2. Subconfluent cells were
dissociated using a cell scraper (30 cm, TPP, Switzerland) and plated
in a flask for 2−3 days.
Detection of Cellular Uptake of the Polymer and Flow
Cytometric Analysis. To detect cellular uptake of polymers, cells
were seeded in a 60 mm dish at a density of 5.0 × 105 cells per dish, in
5 mL of medium. After overnight incubation, the cells were further
incubated for 0.5, 1, 2, or 4 h with either P(NIPAAm-co-CO),
P(NIPAAm-co-CO)-DOPE, P(NIPAAm-co-RH), or P(NIPAAm-co-
RH)-DOPE at 37 or 27 °C in a humidified atmosphere containing 5%
CO2. The concentrations of P(NIPAAm-co-CO) and P(NIPAAm-co-
CO)-DOPE at this point were calculated to be 400 μg/mL based on
the culture medium, whereas those of P(NIPAAm-co-RH) and
P(NIPAAm-co-RH)-DOPE were 200 μg/mL. After incubation,
RAW264.7 cells were washed twice with PBS and harvested with
trypsin/EDTA. Cells were resuspended in PBS, and their cell-
associated fluorescence was detected using a flow cytometer (BD
LSRII Flow Cytometer, BD Biosciences, San Jose, CA, USA).
Fluorescence Microscopy. RAW264.7 cells were seeded in 35
mm glass-bottomed dishes at a density of 2.0 × 105 cells per dish, in 2
mL of medium. After overnight incubation, the cells were further
incubated for 0.5 or 2 h with either P(NIPAAm-co-RH) or
P(NIPAAm-co-RH)-DOPE at 37 or 27 °C in a humidified atmosphere
containing 5% CO2. Following incubation, the RAW264.7 cells were
rinsed twice with PBS and fixed with a 4% paraformaldehyde
phosphate buffer solution for 20 min. The fixed cells were rinsed
with PBS and incubated for 15 min with Hoechst 33258 (5 μg/mL in
PBS) to stain their nuclei. After this time, the cells were then washed
twice with PBS, and the RAW264.7 cells were visualized by
fluorescence microscopy (BZ-9000, Keyence, Osaka, Japan).
RESULTS AND DISCUSSION
■
Synthesis of Fluorescent Polymers. We first synthesized
four fluorescent polymers via radical polymerization. The
B
DOI: 10.1021/acs.biomac.5b00591
Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules
Article
Scheme 1. Syntheses of Fluorescent Polymers
Table 1. Characterization of Synthetic Fluorescent Polymers
a
a
b
polymer
Mn
Mw
Mn/Mw
LCST (°C)
λex (nm)
λem (nm)
P(NIPAAm-co-FL)
P(NIPAAm-co-CO)
P(NIPAAm-co-RH)
P(NIPAAm-co-DA)
26 779
25 360
22 631
21 974
52 994
47 494
45 905
43 596
1.979
1.873
2.028
1.984
30.5
31.5
32.4
31.1
490
376
540
335
515
460
588
526
a
b
Determined by GPC using DMF with 10 mM LiCl. Determined by transmittance measured at 500 nm.
Figure 1. Temperature-responsive fluorescence spectral changes of (A) P(NIPAAm-co-FL) (λex 440 nm), (B) P(NIPAAm-co-CO) (λex 350 nm),
(C) P(NIPAAm-co-RH) (λex 540 nm), and (D) P(NIPAAm-co-DA) (λex 310 nm) in aqueous solution (0.05% w/v) from 25 to 35 °C. Inset:
temperature-dependent optical transmittance for the fluorescent polymers in aqueous solution (500 nm, 0.5% w/v).
synthetic scheme and structures of the target fluorescent
polymers are shown in Scheme 1. For these preparations, we
selected four fluorescent monomers, namely, fluorescein o-
acrylate (FL), 7-(4-trifluoromethyl)coumarin acrylamide (CO),
methacryloxyethyl thiocarbamoyl rhodamine B (RH), and N-
[2-[[[5-(N,N-dimethylamino)-1-naphthalenyl]sulfonyl]amino]-
ethyl]-2-propenamide (DA), which have been previously
verified as being relatively temperature-resistant molecules.
Using AIBN and 3-mercaptopropionic acid as the radical
initiator and chain-transfer agent, respectively, N-isopropyl-
acrylamide (NIPAAm) and the fluorescent monomer (molar
ratio = 1000:1) were copolymerized in anhydrous DMF at 70
C
DOI: 10.1021/acs.biomac.5b00591
Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules
Article
Figure 2. Fluorescence responses of (A) P(NIPAAm-co-FL) (0.05% w/v, λex 440 nm, λem 515 nm), (B) P(NIPAAm-co-CO) (0.05% w/v, λex 350
nm, λem 460 nm), (C) P(NIPAAm-co-RH) (0.05% w/v, λex 540 nm, λem 590 nm), and (D) P(NIPAAm-co-DA) (0.05% w/v, λex 310 nm, λem 500
nm) to pH variation in Britton−Robinson buffer from pH 3−11 at 25−35 °C. Key: Closed circles = 25 °C, open circles = 27 °C, closed triangles =
29 °C, open triangles = 31 °C, closed squares = 33 °C, open squares = 35 °C.
°C for 5 h. The successful introduction of the fluorescent
moiety in the synthetic polymers was confirmed by H NMR
therefore, the fluorescein unit cannot yield strong fluorescence.
Similar phenomena were observed for P(NIPAAm-co-CO) and
P(NIPAAm-co-RH) (Figure 1B,C), as the fluorescent compo-
nents coumarin and rhodamine display the same sensitivity as
that of fluorescein in polar media. In contrast, the presence of
the dansyl moiety in P(NIPAAm-co-DA) resulted in the
opposite characteristics being observed. The fluorescence
intensity of P(NIPAAm-co-DA) was stronger above the
LCST (dehydrated state) than below the LCST (hydrated
state) (Figure 1D). Unlike the other three fluorophores, the
dansyl moiety displays strong emission in nonpolar media, and
this fluorescent property is directly reflected in P(NIPAAm-co-
DA). We could therefore conclude that the on/off emission
responses of these polymers were driven by a combination of
the properties of both PNIPAAm and the fluorescent
molecules. In addition, these properties changed rapidly at
temperatures close to the LCST of the individual polymers.
Furthermore, to confirm that light scattering by the phase-
separated polymer did not cause changes in the fluorescence
intensity, we examined the optical transmittance and
fluorescent spectra of the mixtures of each individual
fluorophore with PNIPAAm. We observed that in all cases
the fluorescence intensity of the mixed solutions remained
largely unchanged, whereas the optical transmittance decreased
above the LCST of PNIPAAm (i.e., 32 °C). These results
confirmed that the increase in the fluorescence intensity of
these fluorescent polymers was caused by the introduction of
the fluorophores to PNIPAAm and not by light scattering
alone.
1
spectroscopy. The number- and weight-averaged molecular
weights (Mn, Mw) were determined by gel permeation
chromatography (GPC). All four polymers were found to
have their LCST at approximately 31 °C and exhibit thermally
reversible solubility changes in response to a temperature
change in water. Detailed physicochemical data for the
prepared synthetic polymers are shown in Table 1. In addition,
as these polymers possess a single terminal carboxyl group, they
have the potential to be further modified.10,18
Characterization of Fluorescent Polymers. We then
chose to investigate the effect of temperature on the fluorescent
behavior of our fluorescent polymers. The spectra of these
polymers exhibited comparable fluorescent behavior to that of
the parent fluorescent monomers (Figure S1). For example, the
absorption and fluorescence maxima for P(NIPAAm-co-FL)
were observed at 444 and 512 nm, respectively, which are
comparable to those of the parent fluorescein (λex = 492 nm,
λem = 514 nm). The temperature-dependent fluorescence
spectra and optical transmittance of P(NIPAAm-co-FL) in
aqueous solution are shown in Figure 1A. The fluorescent
intensity of P(NIPAAm-co-FL) was found to decrease with
increasing temperature, with a sharp change in the intensity of
this polymer being observed close to its LCST (30.5 °C).
Below the LCST, P(NIPAAm-co-FL) assumes its hydrate form
and swollen state driven by the native properties of PNIPAAm.
As fluorescein displays a higher emission quantum yield in polar
media, P(NIPAAm-co-FL) exhibited a strong fluorescence
intensity under such conditions. In contrast, above the LCST,
the polymer assumes its dehydrated globular state and,
We then investigated the influence of pH on the fluorescence
intensity of the synthetic polymers. As previously described, pH
D
DOI: 10.1021/acs.biomac.5b00591
Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules
Article
Figure 3. Effects of incubation temperature on polymer uptake by RAW264.7 cells. (A) P(NIPAAm-co-CO), (B) P(NIPAAm-co-CO)-DOPE, (C)
P(NIPAAm-co-RH), and (D) P(NIPAAm-co-RH)-DOPE. Cells were incubated for 4 h (A, B) or 2 h (C, D) with the polymer at either 37 °C (red)
or 27 °C (blue) in a humidified atmosphere containing 5% CO2.
measurement inside living cells is important for monitoring
cellular internalization pathways, such as receptor−ligand
internalization. Figure 2 shows the fluorescence responses of
the synthetic polymers to both pH (pH 3−11) and temperature
(25−35 °C) in Britton−Robinson buffer. As with temperature
dependence, P(NIPAAm-co-FL) also displayed a response to
pH. The fluorescence intensity increased with basification, and
the polymer showed complete loss of emission at acidic pH
(<5) (Figure 2A). In addition, the fluorescence intensity rapidly
increased between pH 6.0 and 7.0, which is the pH range
corresponding to the pKa of fluorescein (∼6.5).19 It is well-
known that the emission of fluorescein is affected by pH. Under
acidic conditions, fluorescein adopts the lactone conformation
and displays low emission. Although P(NIPAAm-co-CO) has
also been shown to display a pH response (Figure 2B), the
opposite phenomenon was observed with P(NIPAAm-co-FL).
P(NIPAAm-co-CO) exhibited strong fluorescence at low pH
(<8), but it lost its emission at high pH. Comparison of the
properties of CO with P(NIPAAm-co-CO) confirmed that both
displayed comparable pH-dependent fluorescence behavior
(data not shown). These results show that both P(NIPAAm-
co-FL) and P(NIPAAm-co-CO) largely preserved the pH-
responsiveness of their respective fluorophores and were
confirmed to be dual temperature- and pH-responsive
fluorescent polymers. In addition, as with the monomers RH
and DA, the fluorescence intensity of polymers P(NIPAAm-co-
RH) and P(NIPAAm-co-DA) was unaffected by pH.
geometry.20 Figure 3 shows the effect of incubation temper-
ature on the uptake of fluorescent polymers by RAW264.7 cells
measured using flow cytometry. It was observed that cellular
uptake of the fluorescent polymers did not take place below the
LCST but increased drastically above the LCST. The large
increase in cellular uptake of the fluorescent polymers above
the LCST appeared to result from the dehydration of polymer
chains, as observed in our previous studies.15,16 As the
hydrophilic PNIPAAm copolymer becomes hydrated below
the LCST, the hydration layer around the polymer chain
suppresses the interactions between the polymer chains and
between the polymer and the cell membrane. Furthermore,
after dehydration, the polymer chains were able to aggregate
once again by means of hydrophobic interactions between the
chains. This hydrophobic aggregation of polymer chains
resulted in interactions with the cell membranes and thus
cellular uptake was enhanced. Fluorescence microscopy images
of P(NIPAAm-co-RH) and P(NIPAAm-co-RH)-DOPE uptake
into RAW264.7 cells can be seen in Figure 4. The cellular
uptake of fluorescent polymers was not observed below the
LCST, regardless of whether DOPE was present. In addition, it
was found that the LCSTs of P(NIPAAm-co-RH) and
P(NIPAAm-co-RH)-DOPE were comparable (temperature-
dependent optical transmittance curves are shown in Figure
S2). Terminal modification of DOPE did not appear to affect
the hydration layer around the polymer chain below the LCST;
hence, it could be concluded that the fusogenicity of DOPE was
not effective for cellular uptake below the LCST.
Environmental-Dependent Cellular Uptake of Fluo-
rescent Polymers. Following studies into the temperature-
and pH-dependence studies, we chose to study the uptake of
P(NIPAAm-co-CO) and P(NIPAAm-co-RH) into cultured
macrophage cells (RAW 264.7) using both fluorescence
microscopy and flow cytometry (FCM). To enhance cellular
uptake, the fusogenic lipid, L-α-phosphatidylethanolamine,
dioleoyl (DOPE), was conjugated to the fluorescent polymer.
The fusogenicity of DOPE is derived from its tendency to form
a nonlamellar phase (hexagonal phase) due to its cone-shaped
With an incubation time of 2 h, cellular uptake of the
fluorescence polymers was also enhanced above the LCST. In
addition, the DOPE-conjugated polymer was internalized more
rapidly, with an incubation time of only 0.5 h (Figure 4). The
time-dependent intracellular fluorescence intensity of both the
DOPE-conjugated P(NIPAAm-co-CO) and P(NIPAAm-co-
RH) fluorescent polymers, as observed by FCM, is shown in
Figure 5. It can be seen that the modified fluorescent polymers
internalized more rapidly and accumulated to a greater extent
E
DOI: 10.1021/acs.biomac.5b00591
Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules
Article
CONCLUSIONS
■
We conclude that the development of environmentally
responsive fluorescent polymers with four different fluoro-
phores was successful. These PNIPAAm-based fluorescent
copolymers were found to possess two specific characteristics of
interest. First, they possessed sensitive temperature dependence
derived from PNIPAAm and thermally reversible solubility
changes in aqueous solution across the LCST. The polymer
chains were found to display an expanded conformation below
the LCST due to strong hydration and a compact globular form
above the LCST due to dehydration. Second, the properties of
the fluorescent monomers were reflected in the fluorescence
polymers, despite a low NIPAAm-to-fluorescent monomer
molar ratio (1000:1). As FL, CO, and RH have been found to
display high quantum yields in polar solvents, their
corresponding polymers, namely, P(NIPAAm-co-FL), P-
(NIPAAm-co-CO), and P(NIPAAm-co-RH), exhibited strong
emission below their respective LCSTs (i.e., in the hydrated
form). P(NIPAAm-co-FL) and P(NIPAAm-co-CO) were also
found to display pH-dependent fluorescence behavior derived
from the pH-responsive monomers, FL and CO. In cellular
experiments, fluorescence polymers were successfully internal-
ized above the LCST. Cellular uptake was enhanced by the
conjugation of DOPE to the polymers. Studies into the
development of multiresponsive fluorescent polymers and their
applications in cellular imaging are currently underway and will
be reported in due course.
Figure 4. Effects of incubation temperature and polymer modification
with DOPE to polymer uptake by RAW264.7 cells. Incubated with
P(NIPAAm-co-RH) (A, B, E, F) and P(NIPAAm-co-RH)-DOPE (C,
D, G, H). Cells were incubated for 0.5 h (A, C, E, G) or 2 h (B, D, F,
H) with the polymer at 27 °C (A, B, C, D) or 37 °C (E, F, G, H) in a
humidified atmosphere containing 5% CO2.
inside the cells compared with that of the nonconjugated
polymer. These results indicate that DOPE caused a strong
preference for cellular uptake, likely due to its hydrophobicity,
allowing DOPE to be rapidly internalized without the
requirement for a delivery system.
Figure 5. Effects of incubation time on polymer uptake by RAW264.7 cells. (A−D) Flow cytometry peaks of fluorescence intensity in cells treated
with (A) P(NIPAAm-co-CO), (B) P(NIPAAm-co-CO)-DOPE, (C) P(NIPAAm-co-RH), and (D) P(NIPAAm-co-RH)-DOPE. Cells were incubated
with the polymer for 0.5 h (pink), 1 h (yellow), and 2 h (green) at 37 °C in a humidified atmosphere with 5% CO2. (E, F) Summary of mean
fluorescence intensity in cells from flow cytometry over three independent runs.
F
DOI: 10.1021/acs.biomac.5b00591
Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules
Article
ASSOCIATED CONTENT
■
S
* Supporting Information
Absorption and fluorescence spectra, temperature-dependent
1
optical transmittance, and H NMR spectra of fluorescence
polymers. The Supporting Information is available free of
charge on the ACS Publications website at DOI: 10.1021/
acs.biomac.5b00591.
AUTHOR INFORMATION
Corresponding Author
*E-mail: kanazawa-hd@pha.keio.ac.jp.
■
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
Part of this study was supported by a Grant-in-Aid for
Exploratory Research (grant no. 25670081) from the Japan
Society for the Promotion of Science (JSPS), Japan.
REFERENCES
■
(1) Gil, E. S.; Hudson, S. M. Prog. Polym. Sci. 2004, 29, 1173−1222.
(2) Heskins, M.; Guillet, J. E. J. Macromol. Sci., Chem. 1968, 2, 1441−
1455.
(3) Yoshida, R.; Sakai, K.; Okano, T.; Sakurai, Y.; Bae, Y. H.; Kim, S.
W. J. Biomater. Sci., Polym. Ed. 1992, 3, 155−162.
(4) Okano, T.; Yamada, N.; Okuhara, M.; Sakai, H.; Sakurai, Y.
Biomaterials 1995, 16, 297−303.
(5) Kanazawa, H.; Sunamoto, T.; Matsushima, Y.; Kikuchi, A.;
Okano, T. Anal. Chem. 2000, 72, 5961−5966.
(6) Annaka, M.; Tanaka, C.; Nakahira, T.; Sugiyama, M.; Aoyagi, T.;
Okano, T. Macromolecules 2002, 35, 8173−8179.
(7) Uchiyama, S.; Kawai, N.; de Silva, A. P.; Iwai, K. J. Am. Chem. Soc.
2004, 126, 3032−3033.
(8) Gota, C.; Okabe, K.; Funatsu, T.; Harada, Y.; Uchiyama, S. J. Am.
Chem. Soc. 2009, 131, 2766−2767.
(9) Wang, D.; Miyamoto, R.; Shiraishi, Y.; Hirai, T. Langmuir 2009,
25, 13176−13182.
(10) Kobayashi, H.; Nishikawa, M.; Sakamoto, C.; Nishio, T.;
Kanazawa, H.; Okano, T. Anal. Sci. 2009, 25, 1043−1047.
(11) Lowell, B. B.; Spiegelman, B. M. Nature 2000, 404, 652−660.
(12) Karnebogen, M.; Singer, D.; Kallerhoff, M.; Ringert, R.-H.
Thermochim. Acta 1993, 229, 147−155.
(13) Izumi, H.; Torigoe, T.; Ishiguchi, H.; Uramoto, H.; Yoshida, Y.;
Tanabe, M.; Ise, T.; Murakami, T.; Yoshida, T.; Nomoto, M.; Kohno,
K. Cancer Treat. Rev. 2003, 29, 541−549.
(14) Davies, T. A.; Fine, R. E.; Johnson, R. J.; Levesque, C. A.;
Rathbun, W. H.; Seetoo, K. F.; Smith, S. J.; Strohmeier, G.; Volicer, L.;
et al. Biochem. Biophys. Res. Commun. 1993, 194, 537−543.
(15) Hiruta, Y.; Shimamura, M.; Matsuura, M.; Maekawa, Y.;
Funatsu, T.; Suzuki, Y.; Ayano, E.; Okano, T.; Kanazawa, H. ACS
Macro Lett. 2014, 3, 281−285.
(16) Hiruta, Y.; Funatsu, T.; Matsuura, M.; Wang, J.; Ayano, E.;
Kanazawa, H. Sens. Actuators, B 2015, 207, 724−731.
(17) Shea, K. J.; Stoddard, G. J.; Shavelle, D. M.; Wakui, F.; Choate,
R. M. Macromolecules 1990, 23, 4497−4507.
(18) Takei, Y. G.; Aoki, T.; Sanui, K.; Ogata, N.; Okano, T.; Sakurai,
Y. Bioconjugate Chem. 1993, 4, 42−46.
(19) Martin, M. M.; Lindqvist, L. J. Lumin. 1975, 10, 381−390.
(20) Hamai, C.; Yang, T.; Kataoka, S.; Cremer, P. S.; Musser, S. M.
Biophys. J. 2006, 90, 1241−1248.
G
DOI: 10.1021/acs.biomac.5b00591
Biomacromolecules XXXX, XXX, XXX−XXX