A Molecular Chameleon for Mapping Subcellular Polarity in an Unfolded Proteome Environment

Article abstract of DOI:10.1002/anie.201914263

Environmental polarity is an important factor that drives biomolecular interactions to regulate cell function. Herein, a general method of using the fluorogenic probe NTPAN-MI is reported to quantify the subcellular polarity change in response to protein

Full text of DOI:10.1002/anie.201914263

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Accepted Article
Title: A Molecular Chameleon for Mapping Subcellular Polarity of
Unfolded Proteome Environment
Authors: Tze Cin Owyong, Pramod Subedi, Jieru Deng, Elizabeth
Hinde, Jason J Paxman, Jonathan M White, Weisan Chen,
Begona Heras, Wallace W H Wong, and Yuning Hong
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To be cited as: Angew. Chem. Int. Ed. 10.1002/anie.201914263
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10.1002/anie.201914263
Angewandte Chemie International Edition
RESEARCH ARTICLE
A Molecular Chameleon for Mapping Subcellular Polarity of
Unfolded Proteome Environment
Tze Cin Owyong,[a,b] Pramod Subedi,[c] Jieru Deng,[c] Elizabeth Hinde,[d] Jason J. Paxman,[c] Jonathan
M. White,[a] Weisan Chen,[c] Begoña Heras,[c] Wallace W. H. Wong,*[a] and Yuning Hong*[b]
Abstract: Environmental polarity is an important factor in driving
biomolecular interactions to regulate cell function. In this work, a
general method of using the fluorogenic probe NTPAN-MI was
reported to quantify the subcellular polarity change in response to
protein unfolding. NTPAN-MI fluorescence is selectively activated
upon labeling unfolded proteins with exposed thiols to report on the
extent of proteostasis. NTPAN-MI was used to reveal the collapse of
the host proteome caused by influenza A virus infection. The emission
profile of NTPAN-MI contains information of the local polarity of the
unfolded proteome, which can be resolved through spectral phasor
analysis. Under stress conditions that disrupt different checkpoints of
protein quality control, distinct patterns of dielectric constant
distribution in the cytoplasm can be observed. However, in the
nucleus, the unfolded proteome was found to experience a more
hydrophilic environment across all the stress conditions, indicating the
central role of nucleus in the stress response process.
Recent research has shown the connection between misfolded
proteins and the formation of membraneless compartments, such
as stress granules, p-bodies, and nuclear bodies, suggesting the
crucial role of these compartments in buffering proteostasis
stress.[5-7] Although the exact mechanism remains elusive, the
change in the intracellular environment is considered as an
important factor driving the assembly of these dynamic
membraneless condensates.[7,8] A phase separation process
mediates the formation of these bodies where the polarity of the
interior is distinct from the general environment in the cytoplasm
and nucleus.[9] Moreover, the environment in soluble, disordered
aggregates of misfolded proteins has also been reported to be
distinct from those of the ordered, fibrillar structures.[10,11]
Therefore, quantifying the change of environmental polarity could
provide
a
better understanding of the stress response
mechanisms in the cell as well as informing about the
pathogenicity associated with diseases related to protein
misfolding.
Herein, we develop a general strategy to quantify the polarity,
in terms of dielectric constant (e), of the local environment
surrounding unfolded proteins in cells. Our approach is based on
a unique environment-sensitive fluorogenic dye: the fluorescence
is activated upon reaction with unfolded proteins and the emission
profile can be decoded to retrieve quantitative information of the
dielectric constant in the cells. We validate our strategy in vitro by
using purified model proteins and in cells under stress conditions
that promote protein unfolding. We further apply our strategy to
quantify the change of polarity surrounding unfolded proteins in
different intracellular compartments driven by the increase of
unfolded protein load. Compared to our previously reported
approaches to quantifying cytoplasmic unfolded proteins,[12,13] this
multi-responsive probe serves as a “molecular chameleon” that
holds unprecedented advantage for direct visualizing and
quantifying unfolded proteins in both the cytoplasm and the
nucleus, with sensitivity to in situ environmental polarity changes
in different subcellular compartments arising from protein
unfolding.
Introduction
Folding of polypeptide chains into the correct native structure is
essential for proteins to be functional.[1] To maintain the proteome
in the folded state, cells adopt a sophisticated quality control
system, known as proteostasis, to constantly keep the balance of
the synthesis, maturation and degradation of cellular proteins.
This system is governed by a network of over 800 proteins in
humans.[2-3] Environmental perturbations or aberrant proteins
(mutations, translational errors, etc.) can impair the buffering
capacity of proteostasis and result in a backlog of proteins that
are either not fully folded or aggregated, the latter is a common
feature associated with neurodegeneration, cancers and
autoimmune diseases.[4]
[a]
[b]
[c]
[d]
T. C. Owyong, Prof., J. M. White, Dr., W. W. H. Wong
ARC Centre of Excellence in Exciton Science, School of
Chemistry, Bio21 Institute, The University of Melbourne, Parkville,
VIC 3010 Australia
E-mail: wwhwong@unimelb.edu.au
T. C. Owyong, Dr., Y. Hong
Department of Chemistry and Physics, La Trobe Institute for
Molecular Science, La Trobe University, Melbourne, VIC 3086,
Australia
E-mail: Y.Hong@latrobe.edu.au
P. Subedi, J. Deng, Dr., J. J. Paxman, Prof., W. Chen, A/Prof B.
Heras
Results and Discussion
Probe Design Rationale
Our design rationale is to target unfolded proteins in cells using a
cysteine (Cys)-reactive aggregation-induced emission (AIE)
fluorophore. We have recently demonstrated the use of
maleimide (MI) functionalized tetraphenylethene (TPE-MI, 1,
Figure 1A) for measuring the unfolded protein load as a proxy
reporter of the proteostasis capacity inside the cells.[12] The
activation of TPE-MI fluorescence has two criteria: the removal of
the photoinduced electron transfer (PeT) quenching effect of MI
and the restriction of the intramolecular motions of TPE. The MI
Department of Biochemistry and Genetics, La Trobe Institute for
Molecular Science, La Trobe University, Melbourne, VIC 3086,
Australia
Dr., E. Hinde
School of Physics, Department of Biochemistry and Molecular
Biology, Bio21 Institute, The University of Melbourne, VIC 3010,
Australia
Supporting information for this article is given via a link at the end
of the document.
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10.1002/anie.201914263
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RESEARCH ARTICLE
Figure 1. Dye structures and photophysical characterization. (A) Chemical structures of previously reported (1) and newly synthesized thiol-reactive probes (2-4).
(B) HOMO and LUMO energy levels for 4 (NTPAN-MI). (C) Normalized UV-Vis absorbance spectra for compound 1-4 in DMSO. (D) Model thiol-conjugate NTPAN-
MI (4a). (E) Normalized fluorescence spectra of 4a in solvents with different polarity. (F) Correlation of the emission maximum of 4a versus dielectric constant (e) of
solvents. Excitation wavelength: 405 nm. Dye concentration: 50 µM.
moiety enables the specific targeting on Cys residues in the free
thiol form which are usually buried in folded proteins but become
exposed when proteins unfold. Upon reacting with a Cys, the PeT
effect can be suppressed. Meanwhile, the hydrophobic
sidechains on unfolded proteins can provide an environment that
contributes to TPE fluorescence. This unique property enables
the differentiation of Cys residues on unfolded proteins from small
biothiols such as glutathione (GSH) and surface-exposed Cys in
the intracellular redox buffering system.
A copper catalyzed amination of aryl-bromide cores led to amine
analogs that were further converted to maleimide in the final
products using maleic anhydride. The final products 2-4 were
characterized by 1H, 13C NMR, high-resolution mass spectrometry,
and FT-IR spectroscopy (refer to SI). The structures were further
confirmed by single crystal X-ray crystallography (Figure S42 and
Table S3 – S5). The core of triphenylacrylonitrile (TPAN) adopts
a highly twisted geometry with the torsion angles ranging from
121.61 – 146.74° between the phenyl rings and the central double
bond (Figure S43). Such geometry reduces aggregation-caused
quenching from strong p-p stacking (Figure S44-46).[15,17]
Comparison of the UV-Vis absorption spectra of dyes revealed
the effect of the electron donating groups on the p conjugate
system of TPAN (Figure 1C). There are red-shifts in the
absorption maximum from 2 (318 nm) to methoxy-substituted 3
(350 nm) and dimethylamino-substituted 4 (393 nm). With the
largest red-shift and strongest donor-acceptor effect, dye 4
(NTPAN-MI) was selected for subsequent studies.
To achieve the environmental sensitivity, our strategy involved
replacing the TPE unit with a push-pull AIE fluorophore (Figure
1A). Push-pull dyes with both electron donor and acceptor
experience excited-state charge transfer, which is stabilized by
interaction with the dipoles of solvent and resulting in emission
red-shift in more polar solvents.[13] Being sensitive to polarity,
push-pull dyes have been extensively used to study the
biophysical properties of lipid membranes and for the detection of
biomolecular interactions such as protein-lipid interaction.[14] On
the other hand, the activation of AIE dyes through restriction of
intramolecular motions has been used in designing AIE probes for
protein detection and characterization.[15] To construct a push-pull
AIE fluorophore while retaining a similar molecular structure, we
changed one of the phenyl rings of TPE to an electron-
withdrawing cyano group (Figure 1A).[16] Methoxy and
dimethylamino substituents are used as electron donors, which
also improve the compatibility of the dyes in aqueous media.[13]
NTPAN-MI is inherently non-fluorescent until it reacts with a
thiol via the C=C on the MI.[18] This observation was supported by
DFT calculations showing a low-lying np* state between HOMO
and LUMO of NTPAN induced by the MI group (Figure 1B). This
energy level that contributed to the non-radiative decay of excited
NTPAN-MI was not present in the saturated succinimide analog
which was formed after thiol conjugation to the MI (Figure S1). In
addition, we observed that electron density for HOMO levels were
more localized around the electron-donating groups while the
LUMO were closer to the electron-withdrawing cyano group
(Figure 1B). Such electron distribution is characteristic of push-
pull dyes with photoinduced charge transfer, which results in the
solvatochromism in their fluorescence.[13] In order to study the
Dye Synthesis and Photophysical Characterization
The synthetic routes to compound 2-4 are shown in Scheme S1
in the supporting information. In short,
a
Knoevenagel
condensation reaction between respective diarylketones and
bromobenzyl cyanide gave the donor-acceptor cores of interest.
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Figure 2. In vitro fluorescence studies with model proteins. (A) Fluorescence kinetic traces of NTPAN-MI with folded/unfolded β-lac and GSH. Denaturant: 6 M urea
in PBS; protein concentration: 250 µM; dye concentration: 50 µM. Excitation: 405 nm; emission: 540 nm. (B) Crystal structures of DsbA and DsbL, adapted from
Ref 19. The red box highlights the position of the surface-exposed Cys on the proteins. (C) Emission spectra of NTPAN-MI with oxidized (Ox) or reduced (Red)
forms of DsbA and DsbL in folded/unfolded (Urea) status. Sample preparation showed in SI. (D) Endpoint intensity of NTPAN-MI fluorescence with folded
oxidized/reduced DsbL, folded reduced DsbA in 1:1 or 1:2 protein-to-dye molar ratio, or with unfolded reduced DsbA. Data extracted from kinetic traces shown in
Figure S7B. Protein concentration: 25 µM; dye concentration: 25 or 50 µM; excitation and emission filter: 390/20 and 540/50 nm respectively. n =3 biological
replicates; mean ± s.d.
fluorescence behavior, the thiol-bound form of NTPAN-MI (4a)
was prepared (Figure 1D and Scheme S1). The UV-Vis
absorption spectra of 4a exhibited negligible shift among solvents
with different polarity (Figure S2-3). The emission spectra
(excitation at 405 nm), on the other hand, displayed a large red-
shift with the increase of solvent polarity. The emission peak
shifted from 463 nm in nonpolar toluene to 586 nm in phosphate
buffered saline (PBS) solution (Figure 1E). The correlation
between the emission maximum and the dielectric constant of the
solvent is in a linear relationship (Figure 1F). The nature of the
absorbance and emission profiles of the thiol-bound NTPAN-MI
would allow us to estimate and report on the local environment of
bound unfolded proteins in a cell.
unfolded protein, and there was minimal self-aggregation of the
GSH-bound NTPAN-MI.
Next, the role of hydrophobic surface in NTPAN-MI reactivity
was investigated with two thiol disulfide oxidoreductases (Dsb)
which contain
a surface-exposed Cys residue in distinct
environments. DsbA and DsbL proteins share a very similar three-
dimensional structure encompassing two cysteines, one buried
and one surface exposed, that could be oxidized to disulfide
bonded or reduced to two free thiols.[19] A major difference
between these proteins is the local environment surrounding the
surface exposed cysteine: DsbA is hydrophobic and DsbL,
hydrophilic (Figure 2B).[19] We first tested the Dsb proteins in their
oxidized disulfide bonded form, which, as expected, could not
activate the NTPAN-MI fluorescence (Figure 2C). Incubation of
NTPAN-MI with reduced Dsb proteins, yielded approximately 25-
fold fluorescence enhancement on DsbA but minimal
enhancement on DsbL, indicating the interaction between
Surface Exposed vs. Buried Cys on Dye Reactivity
To validate this hypothesis, the reactivity of NTPAN-MI was first
tested in vitro with model proteins. β-lactoglobulin (β-lac) with 5
buried Cys in the thiol form was chosen as a test protein. NTPAN-
MI showed weak fluorescence in the presence of folded β-lac but
became highly emissive when β-lac was unfolded in 6 M urea
(Figure 2A). The kinetics curve indicated NTPAN-MI is able to
react with exposed Cys rapidly when β-lac unfolds. Addition of
thiol-blocking reagents, such as N-methylmaleimide (NMM),
reduced NTPAN-MI emission intensity effectively, especially with
unfolded proteins, indicating they are competing for the same
species, i.e. Cys on the proteins (Figure S4). Replacing Cys-
reactive MI to an inert Br substituent yielded a control probe,
NTPAN-Br, which exhibited high background fluorescence in
aqueous solution (Figure S5). No fluorescence changes could be
observed between folded and unfolded proteins, further
confirming the importance of the MI moiety in the detection
process. On the other hand, the presence of hydrophilic GSH,
even in much higher concentration, could not induce NTPAN
fluorescence to the same extent as the unfolded β-lac (Figure S6).
These results suggest that the reactivity of NTPAN-MI with
hydrophilic GSH is much lower than that with thiol containing
NTPAN-MI and Cys is more favored in
a hydrophobic
environment. When protein-to-dye molar ratios were changed
from 1:1 to 1:2, the fluorescence intensity remained comparable,
implying that NTPAN-MI does not perturb the folded structure of
the proteins or access the buried Cys residue (Figure 2D). The
conjugation of one NTPAN-MI molecule with folded DsbA was
further confirmed by intact protein mass spectrometry (Figure S7).
This fluorescence intensity for the hydrophobic DsbA, however,
was negligible compared to the significant 120-fold increase in
intensity for reduced DsbA in the unfolded state (25 fold). This
suggests that when reacted with surface-exposed Cys, the dye
molecule is not fully restricted and the fluorescence remains
relatively weak, when compared to the substantial fluorescence
emission of unfolded DsbA conjugated NTPAN-MI.
Cys reactivity is dependent on pH.[20] To test the influence of
pH on NTPAN-MI reactivity, we measured the fluorescence at the
physiological pH range (~4-8) (Figure S8-S9). Minimal change in
fluorescence intensity was observed in the presence of folded
reduced DsbA with NTPAN-MI between pH 4.27-7.10 with a slight
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Figure 3. Intracellular distribution of unfolded proteins revealed by NTPAN-MI. (A) Confocal images of control and stressed HeLa cells stained with NTPAN-MI with
ER-TrackerTM Red and DRAQ5TM counterstain to view colocalization with endoplasmic reticulum (ER) and nucleus respectively. PER and PNUC are the calculated
Pearson correlation values for NTPAN-MI in ER and nucleus respectively. Scale bar: 20 µm. (B) Enlarged images of insets in panel A. Scale bar 20µm.
increase at pH 7.89 (Figure S9A). Normalized fluorescence
kinetic traces did not show any significant deviation between the
varying pH, implying that the reactivity of NTPAN-MI with Cys is
less likely to be influenced by pH variation within the physiological
range (Figure S9B).
proteostasis stress. Puromycin, tunicamycin, brefeldin A, and
MG132, which interfere with the protein synthesis, folding,
transport and degradation pathways, respectively, were chosen
in the study.[21-24] Disruption of any of these pathways could lead
to the backlog of premature or unfolded proteins accumulated in
the cells, which was confirmed by the significant increase of
NTPAN-MI fluorescence of prelabeled cell protein lysates (Figure
S12). The fluorescence was normalized to total protein
concentration under each condition to avoid the variation of
protein concentration caused by the stressors. The fluorescence
enhancement in stressed cells could also be seen from confocal
images (Figure 3). The NTPAN-MI staining patterns in the
cytoplasm were highly colocalized with ER-TrackerTM Red, with
the Pearson correlation factor (PER) between 0.85-0.91 under all
conditions. Despite of penetration of NTPAN-MI into the nucleus,
we obtained negative correlation values (PNUC) with the DNA stain
DRAQ5TM, suggesting that proteins rather than DNA were being
labelled by the dye. Distinct protein condensates with bright
NTPAN-MI fluorescence were observed in the nucleus under
certain stress conditions (Figure 3B), in particular, puromycin
treatment, indicating the unfolded protein load also increased in
the nucleus in response to such proteostasis stress.
Visualizing Unfolded Proteins in Cytoplasm and Nucleus
The in-vitro experiments with model proteins confirmed that the
major fluorescence enhancement of NTPAN-MI could be
attributed to the reaction with unfolded proteins containing non-
disulfide bonded Cys residues. Next, the behavior of NTPAN-MI
in cells was examined. Cytotoxicity assay showed that NTPAN-
MI posed no significant toxicity towards cells at concentration up
to 100 µM and incubation time as long as 12 h (Figure S10).
Confocal imaging revealed that NTPAN-MI was readily
permeable to not only the plasma membrane but also the nuclear
membrane after 30 min of staining (Figure 3). To examine the
intracellular distribution of NTPAN-MI (pseudo green), the cells
were counterstained with ER-TrackerTM Red (pseudo red) and
DRAQ5TM (pseudo blue) to localize the ER and nucleus
respectively. The settings of the three channels were optimized to
avoid signal bleed-through (Figure S11)
In control/untreated cells, NTPAN-MI displayed dim
fluorescence signals homogeneously distributed in the cells
(Figure 3). The ability of NTPAN-MI to detect potentially unfolded
proteins in cellulo was assessed using the drugs that induce
Mapping Subcellular Polarity of the Unfolded Proteome
Environment
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Figure 4. Mapping and quantifying intracellular polarity of unfolded protein environment. (A) Representative images of intracellular dielectric constant (e) distribution
in untreated and stressed cells stained by NTPAN-MI resolved from spectral phasor analysis. Excitation: 405 nm; emission: acquired from 415-700 nm with 8.9 nm
bin widths to obtain 32 channels. Scale bar: 20 µm. (B) Changes in e distributions of unfolded protein environment in ER, nucleus and cytosol in brefeldin A, MG132,
tunicamycin, and puromycin treated cells compared to untreated cells. A positive or negative change in the plots reflects a respective increase or decrease in the
pixel population within that specific e range. The analysis is based on 10 images with total 38-56 cells analyzed in each condition.
Following the characterization of dye distribution in cells, we took
advantage of the solvatochromism of NTPAN-MI (cf. Figure 1E,
Figure S3) to assess the local polarity of the bound proteins in
cells. Spectral phasor analysis is a powerful method that allows
us to deconvolute and identify the emission spectra of pixels on a
global scale in an acquired image.[25,26] Upon excited by 405 nm
laser, confocal images were collected in every 8.9 nm from 415 to
700 nm in 32 channels (Figure S13). We then focused on using
the phasor angle to determine emission spectral wavelength from
the 32 channel images to estimate the dielectric constant (e) of
individual pixels. By using spectral phasor analysis, we found the
e values varied in different compartments of the cells. The labeled
unfolded proteins experienced e in a range of 22-32 in the ER, 32-
36 at the interface of ER and cytoplasm, 36-45 inside the nucleus,
and 45-50 on the plasma membrane in the control/untreated cells.
These values are generally consistent with the dielectric constant
in cytoplasm and nucleus measured by electrochemical
methods.[27,28]
addition of the stressors puromycin, tunicamycin, brefeldin A and
MG132, the overall intracellular environment became even more
discrete (Figure 4A). Although each stressor led to a distinct
pattern of e distribution, the common feature was that the unfolded
protein environment in the nucleus became much more
hydrophilic (e shifting to 45-54).
To further quantify the intracellular e distribution in a larger
population of cells, we separated NTPAN-MI signals into three
groups based on subcellular locations and fluorescence intensity,
namely the ER, nucleus and cytosol (Figure S15). 10 images with
total 38-56 cells in each condition were acquired for the spectral
phasor analysis. The same spectral gating strategy was applied
to the estimation and subsequent quantification of environmental
polarity in individual cellular compartments in all acquired images
(Figure S15, Table S2). Figure 4B shows the comparison of the
population of pixels at a specific dielectric constant range of the
stressed cells when compared to those in control cells. A positive
or negative change in the plots reflects a respective increase or
decrease in the pixel population within that specific polarity range.
A positive shift to lower e region was found in the ER when the
cells were treated with ER stressors such as brefeldin A and
tunicamycin (Figure 4B). Brefeldin A blocks protein transport from
ER to Golgi, while tunicamycin inhibits the N-linked glycosylation
of proteins in ER.[21,22] Both lead to the build-up of misfolded
proteins in the ER, possibly with increased hydrophobic patches
exposed, resulting in a more hydrophobic environment for
NTPAN-MI. MG132, a proteasome inhibitor, causes a backlog of
ubiquitylated proteins that would normally be degraded in the
cells.[24] Spectral profile of NTPAN-MI reflected that these proteins
in both ER and cytosol were in a more hydrophilic environment
upon MG132 treatment. This was in line with previously reported
observation that ubiquitylated proteins in MG132-treated cells
were more soluble in PBS, indicating that these proteins were
The above results demonstrated that the unfolded proteins in
the ER are mostly in a hydrophobic environment with low e while
in the nucleus the environment is more hydrophilic. Charged
species are most likely to contribute to the dielectric properties of
the intracellular environment.[28] The distinct e observed in the
nucleus could be attributed to the environment rich in highly
charged nucleic acids. The interaction of the unfolded proteins
with nucleic acids creates an environment with dielectric property
differing from either non-charged protein-rich (lower e) or aqueous
environment (higher e). This hypothesis is also supported by the
previous reported e value of stress granules, composed of
proteins and RNAs, sitting in the range of acetonitrile and DMSO
(e = 40-50).[9,28] As a control experiment, permeating the cell
membrane with Triton X-100 resulted in
a highly polar
environment in all cell compartments (Figure S14). Upon the
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10.1002/anie.201914263
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RESEARCH ARTICLE
more likely to reside in polar environments.[29] Puromycin, a
protein synthesis inhibitor, did not induce noticeable change in the
unfolded protein environment in the cytoplasm. However, across
all stress treatments, the nucleus had a large shift in the
population of pixels in the more hydrophilic, higher e region. Such
changes could arise from the impairment of nucleocytoplasmic
transport pathways under stress.[30,31] Recent literature also
pointed out the interaction of unfolded proteins and RNA in the
nucleus as a strategy to temporarily store the dynamic unfolded
proteins and prevent the formation of irreversible protein
aggregates.[32] This is an exciting observation with further
experiments currently being carried out.
Measuring Proteostasis Collapse in Influenza Virus Infected
Cells
With the encouraging results in drug-induced stress models,
NTPAN-MI was further applied to assess proteostasis stress
induced by virus infection. Cells with successful influenza A virus
(IAV) infection were recognized by the immunostaining of IAV
nucleoproteins (NP). Flow cytometry analysis revealed
a
significant increase in NTPAN-MI fluorescence in the infected
(IAV+) group, in comparison to the uninfected (IAV-) group, in
both suspension P815 and adherent A549 cell-lines (Figure 5A,B;
Figure S16). Confocal imaging showed that the NTPAN-MI
staining pattern changed from homogenous distribution in
uninfected cells to discrete puncta in IAV-infected cells (Figure
Figure 5. NTPAN-MI reveals proteostasis collapse in influenza virus infected
cells. (A) P815 and (B) A549 cells were mock-infected or infected with IAV (PR8
(H1N1)) at MOI=5 for overnight. Cells were then stained with NTPAN-MI (50
µM) for 30 min and analysed by flow cytometry. Two independent experiments
were conducted with similar results, and each independent experiment included
3 biological replicates; mean ± s.d. **P < 0.01; ****P < 0.0001. (C) Confocal
images of NTPAN-MI stained A549 cells with anti-IAV NP counterstain. Scale
bar: 20 µm.
5C).
A similar phenomenon has been observed on the
the pathogenesis, finding biomarkers and screening drugs that
rectify proteostasis for the treatment of neurodegenerative
diseases.
autophagosome marker LC3 protein upon lipidation in cells with
IAV infection.[33] Interestingly, little colocalization was observed of
NTPAN-MI fluorescence with NP immunofluorescence, possibly
due to the Cys residues being buried in NP [Figure 5C, S17B (Z-
stack)].[34] The IAV virus hijacks the host proteostasis machinery
to facilitate the synthesis and processing of viral proteins for their
own replication.[35] Our results showed that such process disrupts
the protein quality control of the host cells, as revealed by the
increase of NTPAN-MI fluorescence. Spectral phasor analysis,
however, was not applied to this particular case due to the
requirements of cell permeabilization for NP immunostaining (cf.
Figure S14).
Acknowledgements
We thank Dr Peter Lock and LIMS Bioimaging Platform, La Trobe
University for the technical support and access to the confocal
microscope and flow cytometer. We thank Dr Swati Varshney,
A/Prof Nicholas A. Williamson and Bio21 Mass Spectrometry and
Proteomics Facility for technical support and access to mass
spectrometers. This work was supported by grants to Y.H.
(Australian Research Council DE170100058, Rebecca L. Cooper
Medical Research Foundation PG2018043, National Health and
Medical Research Council GNT1161803 and Australia-China
Science and Research Fund-Joint Research Centre on Personal
Health Technologies), WWHW (Australian Research Council
CE17100026) and JMW (Australian Synchrotron for beamtime via
the Collaborative Access Program Project Number 13618b).
Conclusion
In summary, a simple approach to quantifying the change of
intracellular polarity accompanying protein misfolding was
presented. NTPAN-MI fluorescence can report on the unfolded
protein load not only in the cytoplasm but, for the first time, in the
nucleus. Spectral phasor analysis of protein-bound NTPAN-MI
solvatochromism reveals the crosstalk between the nuclear and
cytoplasmic protein quality control, an area still underexplored.
Our method provides possibilities to investigate the role of
subcellular polarity in driving fundamental biological processes
involved in protein quality control and stress response. This
method would find many applications including in understanding
Keywords: Fluorescent probes • solvatochromism •
aggregation-induced emission • unfolded protein • proteostasis
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This article is protected by copyright. All rights reserved.

10.1002/anie.201914263
Angewandte Chemie International Edition
RESEARCH ARTICLE
Entry for the Table of Contents
RESEARCH ARTICLE
Here we report a strategy based on a
multi-responsive probe, in
combination with spectral phasor
analysis, to monitor the change of
subcellular polarity in response to
protein unfolding.
Tze Cin Owyong, Pramod Subedi, Jieru
Deng, Elizabeth Hinde, Jason J.
Paxman, Jonathan M. White, Weisan
Chen, Begoña Heras, Wallace W. H.
Wong,* Yuning Hong*
Page No. – Page No.
A Molecular Chameleon for Mapping
Subcellular Polarity of Unfolded
Proteome Environment
This article is protected by copyright. All rights reserved.