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Journal ArticleDOI

A membrane-activatable near-infrared fluorescent probe with ultra-photostability for mitochondrial membrane potentials

07 Jun 2016-Analyst (The Royal Society of Chemistry)-Vol. 141, Iss: 12, pp 3679-3685

TL;DR: A photostable near-infrared (NIR) fluorescent dye for monitoring MMP, named NIMAP, is non-fluorescent in aqueous solution and can be activated by cell membranes, providing high fluorescence contrast and low background fluorescence.

AbstractMitochondrial membrane potential (MMP) is a frequently used indicator for mitochondrial function Herein, we report a photostable near-infrared (NIR) fluorescent dye for monitoring MMP This new probe, named NIMAP, is non-fluorescent in aqueous solution and can be activated by cell membranes, providing high fluorescence contrast and low background fluorescence NIMAP has been validated for monitoring MMP in living mammalian cells and in mice Due to the large fluorescence response, low fluorescence background, high photostability, and excellent tissue penetration resulting from red-shifted excitation and emission in the "optical window" above 600 nm, broad applications of this new probe are expected

Summary (2 min read)

Introduction

  • Mitochondrial membrane potential (MMP) is required for cellular respiration and ATP synthesis.
  • Measuring absolute MMP in living cells is technically challenging.
  • 11 †Electronic supplementary information (ESI) available: Procedures for chemical synthesis, spectroscopic characterization, experimental optimization and comparison, and time-lapse movies.
  • NIMAP is highly attractive, because not only does it command the above-mentioned advantages of a NIR probe, but it also shows a large fluorescence response to MMP changes, membrane-activated fluorescence and minimal background signals, ultra-photostability, and excellent tissue penetration in live animals.

Materials and general methods

  • All chemicals and reagents were purchased from Fisher Scientific (Pittsburgh, PA) or Sigma-Aldrich (St. Louis, Mo) unless specified elsewhere.
  • Plasmid DNA was purified using Syd Laboratories Miniprep columns (Malden, MA).
  • The structure and purity of NIMAP were analyzed with ESI-MS, 1H NMR, and 13C NMR.
  • ESI-MS was run on an Agilent LC-TOF system by direct fusion.
  • NMR spectra were recorded on a Varian Inova 400 instrument with chemical shifts relative to tetramethylsilane.

Spectroscopic characterization

  • A monochromator-based Synergy Mx Microplate Reader (BioTek, Winooski, VT) was used to record all spectra.
  • To measure the fluorescence excitation and emission spectra of NIMAP, the authors prepared NIMAP in liposomes as described elswhere.21 L-α-Lecithin egg yolk and cholesterol were purchased from EMD Millipore (Billerica, MA) and Alfa Aesar (Ward Hill, MA), respectively.
  • The resulting mixture was then sonicated 5 min on a probe sonicator to yield a slightly hazy and transparent solution.
  • To record the excitation spectrum, the emission wavelength was set at 730 nm, and the excitation scanned from 500 nm to 700 nm.
  • Mammalian cell culture and dye loading Human embryonic kidney (HEK) 293T cells or cervical cancer HeLa cells were maintained in T25 flasks containing 5 mL Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and incubated at 37 °C with 5% CO2 in humidified air.

Mammalian cell transfection

  • The prepared transfection mixture was then added to cells and incubated for 2 h at 37 °C.
  • Next, pre-warmed, fresh DMEM containing 10% FBS was used to replace the transfection medium.
  • On the following day, cells were stained with NIMAP before imaging.

Fluorescence imaging

  • Fluorescence microscopy was performed with a Leica SP5 inverted confocal fluorescence microscope with the spectral imaging capability (Leica, Boston, MA), unless otherwise stated.
  • A 40× water lens was used for all imaging studies.
  • To record the emission spectrum for NIMAP loaded into live-cell mitochondria, the emission bandwidth was set at 10 nm and images were taken from 650 nm to 790 nm.

Designing and preparation of NIMAP

  • The authors development of the new NIR MMP probes originated from a Förster resonance energy transfer (FRET) experiment in their laboratory, which used a QSY-21 core structure as a dark quencher.
  • The authors prepared a QSY-21 labeled biotin derivative (2), and observed mitochondria-like organellar fluorescence in cultured mammalian cells stained with 100 μM of the dye (ESI Fig. S1A†).
  • Optimization of NIMAP loading into mammalian cells.
  • By comparison, the background fluorescence of HEK 293T cells stained with a conventional MMP dye, Rhodamine 123, was strong before washing off excess dye molecules (Fig. S3†).

Mitochondrial localization of NIMAP

  • To further confirm the mitochondrial localization of NIMAP, the authors loaded the molecule into HEK 293T cells simultaneously expressing a green fluorescent mWasabi protein fused to a mitochondrial localization tag (Mito-mWasabi).
  • A previous molecular dynamics (MD) study suggests that QSY-21 fluorescence is quenched by ring rotations and electron transfer in the excited state.
  • The authors also measured the absorbance of NIMAP in an aqueous solution, which was similar to the profile of NIMAP fluorescence excitation in liposomes (Fig. 4).
  • The authors observed much weaker signals for Rhodamine 123-stained cells, compared to NIMAP-stained cells; and almost no signal for the other two groups.

Conclusions

  • In summary, starting from a nonfluorescent dark quencher QSY-21, the authors have developed a new NIR probe, NIMAP, for monitoring MMP in living mammalian cells and in vivo.
  • Lipid membranes can activate the fluorescence of NIMAP, and therefore the loading procedures can be simplified to gain low fluorescence background and high fluorescence contrast.
  • When NIMAP was loaded into the mitochondria of living mammalian cells, it responded to MMP changes induced by chemicals with high sensitivity.
  • Because of its excitation and emission at wavelengths above 600 nm, excellent penetration of mouse skin tissue was also observed.
  • In addition, the mitochondrial localization capability of the QSY-21 scaffold may further be exploited to construct other mitochondriatargeting fluorescent probes or selectively deliver inhibitors of mitochondrial enzymes.

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UC Riverside
UC Riverside Previously Published Works
Title
A membrane-activatable near-infrared fluorescent probe with ultra-photostability for
mitochondrial membrane potentials.
Permalink
https://escholarship.org/uc/item/7gg0k6sp
Journal
The Analyst, 141(12)
ISSN
0003-2654
Authors
Ren, Wei
Ji, Ao
Karmach, Omran
et al.
Publication Date
2016-06-01
DOI
10.1039/c5an01860a
Peer reviewed
eScholarship.org Powered by the California Digital Library
University of California

Analyst
PAPER
Cite this: Analyst, 2016, 141, 3679
Received 9th September 2015,
Accepted 28th October 2015
DOI: 10.1039/c5an01860a
www.rsc.o rg/analys t
A membrane-activatable near-infrared uorescent
probe with ultra-photostability for mitochondrial
membrane potentials
Wei Ren,
a
Ao Ji,
a
Omran Karmach,
b
David G. Carter,
c
Manuela M. Martins-Green*
b
and Hui-wang Ai*
a
Mitochondrial membrane potential (MMP) is a frequently used indicator for mitochondrial function.
Herein, we report a photostable near-infrared (NIR) uorescent dye for monitoring MMP. This new probe,
named NIMAP, is non-uorescent in aqueous solution and can be activated by cell membranes, providing
high uorescence contrast and low background uorescence. NIMAP has been validated for monitoring
MMP in living mammalian cells and in mice. Due to the large uorescence response, low uorescence
background, high photostability, and excellent tissue penetration resulting from red-shifted excitation and
emission in the optical window above 600 nm, broad applications of this new probe are expected.
Introduction
Mitochondrial membrane potential (MMP) is required for cel-
lular respiration and ATP synthesis.
1
It is also important for
intracellular calcium dynamics, the production of reactive
oxygen species, neural synapse, inflammation, cell prolifer-
ation, and cell death.
26
A direct monitoring of MMP is, there-
fore, of high interest. Previously, tetraphenylphosphonium
(TPP
+
)-sensitive microelectrodes have been used to measure
the membrane potentials of isolated mitochondria or mito-
chondria in permeabilized cells.
7
Measuring absolute MMP in
living cells is technically challenging.
8
There are only few
reports, which used end-point measurements based on radio-
isotope tracers to assay absolute MMP in suspensions of living
cells.
9,10
On the other hand, fluorescent MMP sensors are
popular for assessing MMP in living cells,
11
despite that only
qualitative or semi-quantitative information can typically be
derived from those imaging experiments. Fluorescent MMP
dyes are mostly lipophilic cationic compounds. When applied
to healthy cells, they tend to accumulate in the inner mito-
chondrial inner membrane, because healthy cells maintain
cytoplasmic and mitochondrial transmembrane potentials to
induce a negatively charged mitochondrial inner membrane.
12
The distribution of these probes is aected by the transmem-
brane electric fields, following the Nernst equation.
13
There-
fore, the observed mitochondrial fluorescence intensity can be
utilized as an indicator for MMP, although other factors, such
as the cytoplasmic transmembrane potential, the volume ratio
of the mitochondria and a whole cell, and the participation
coecients of dyes in mitochondria and the cytosol, also have
an impact on the mitochondrial accumulation of these
cationic molecules.
Fluorescent probes with excitation and emission in the
600900 nm spectral region are desirable for mammalian
studies, due to the weak absorbance of hemoglobin, myoglo-
bin, melanin, and water in this NIR optical window.
14,15
More-
over, compared to light at shorter wavelengths, light with
wavelengths matching this optical window overlaps less with
the excitation of endogenous chromophores, such as flavins
and NADH.
16
These lower-energy photons also cause less
photodamage to living cells and tissues.
17
Not surprisingly, a
current trend is to develop novel fluorescent dyes and sensors
that fluoresce in this NIR spectral region, such as recently
reported NIR dyes that are sensitive to cytoplasmic membrane
potentials or that can highlight the mitochondrial structure in
live cells.
18,19
Common MMP dyes,
11
such as TMRM (tetra-
methylrhodamine methyl ester), TMRE (tetramethylrhodamine
ethyl ester), Rhodamine 123, and JC-1, and a recent TPE-indo
dye
20
all have fluorescence excitation and emission peaks less
than 600 nm. Other concerns of these existing MMP probes
include nonspecific background fluorescence, poor locali-
zation, high cytotoxicity, and unsatisfactory photostability.
11
Electronic supplementary information (ESI) available: Procedures for chemical
synthesis, spectroscopic characterization, experimental optimization and com-
parison, and time-lapse movies. See DOI: 10.1039/c5an01860a
These two authors contributed equally to this work.
a
Department of Chemistry, University of California, Riverside, CA 92521, USA.
E-mail: huiwang.ai@ucr.edu
b
Department of Cell Biology and Neuroscience, University of California, Riverside,
CA 92521, USA. E-mail: manuela.martins@ucr.edu
c
Institute for Integrative Genome Biology, University of California, Riverside,
CA 92521, USA
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Herein, we report the development of a NIR probe (designated
NIMAP) for monitoring MMP in living mammalian cells and
mice. NIMAP is highly attractive, because not only does it
command the above-mentioned advantages of a NIR probe,
but it also shows a large fluorescence response to MMP
changes, membrane-activated fluorescence and minimal
background signals, ultra-photostability, and excellent tissue
penetration in live animals.
Experimental
Materials and general methods
All chemicals and reagents were purchased from Fisher Scien-
tific (Pittsburgh, PA) or Sigma-Aldrich (St. Louis, Mo) unless
specified elsewhere. Plasmid DNA was purified using Syd Labo-
ratories Miniprep columns (Malden, MA). The scheme and
procedure to synthesize NIMAP are outlined in the ESI. The
structure and purity of NIMAP were analyzed with ESI-MS,
1
H NMR, and
13
C NMR. ESI-MS was run on an Agilent LC-TOF
system by direct fusion. NMR spectra were recorded on a
Varian Inova 400 instrument with chemical shifts relative to
tetramethylsilane. Stock solutions of NIMAP were made in
DMSO (Sigma-Aldrich).
Spectroscopic characterization
A monochromator-based Synergy Mx Microplate Reader
(BioTek, Winooski, VT) was used to record all spectra. The
absorption spectrum of NIMAP was measured by preparing
NIMAP (50 μM) in a 1 : 1 (v/v) mixture of methanol and Tris-
HCl aqueous buer (50 mM, pH 7.4). Absorbance values of
NIMAP at 600 nm was measured at various pH conditions by
preparing a series of 1 : 1 (v/v) mixtures of methanol and
aqueous solutions containing citric acid (200 mM) and phos-
phate (200 mM) with a pH range from 3 to 11. To measure the
fluorescence excitation and emission spectra of NIMAP, we
prepared NIMAP in liposomes as described elswhere.
21
L-α-Lecithin egg yolk and cholesterol were purchased from
EMD Millipore (Billerica, MA) and Alfa Aesar (Ward Hill, MA),
respectively. Cholesterol (2.5 mg) and
L-α-Lecithin egg yolk
(10 mg) in chloroform : methanol (v/v 3 : 2) were vacuumed on
a rotary evaporator for 3 h to form a thin, oily film on the glass
wall of the flask. Next, double distilled water (ddH
2
O, 1 mL)
was added and the rotation of the rotary evaporator continued
for another 15 min to form an oil/water mixture in the flask.
The resulting mixture was then sonicated 5 min on a probe
sonicator to yield a slightly hazy and transparent solution.
Formed liposomes were next diluted with equal volume
phosphate buered saline (PBS, pH 7.4). NIMAP was added to
a final concentration of 100 μM. The mixture was incubated at
room temperature for 15 min. To record the emission
spectrum, the excitation wavelength was set at 630 nm, and
the emission scanned from 650 nm to 870 nm. To record the
excitation spectrum, the emission wavelength was set at
730 nm, and the excitation scanned from 500 nm to 700 nm.
To determine the responsiveness of NIMAP fluorescence to pH
changes, we mixed NIMAP (50 μM) with a series of 9 : 1 (v/v)
mixtures of glycerol and the aforementioned aqueous solu-
tions with a pH range from 3 to 11. Fluorescence at 710 nm
was recorded with the excitation wavelength set at 650 nm.
Mammalian cell culture and dye loading
Human embryonic kidney (HEK) 293T cells or cervical cancer
HeLa cells were maintained in T25 flasks containing 5 mL
Dulbeccos Modified Eagles Medium (DMEM) supplemented
with 10% fetal bovine serum (FBS) and incubated at 37 °C
with 5% CO2 in humidified air. Cells at 80% confluence were
passaged into 35 mm culture dishes in a ratio of 1 : 5 or 1 : 20
for transfection or direct dye loading on the following day.
To stain cells, fluorescent dyes were added into the cell culture
medium, and the incubation condition was typically 45 min
at 37 °C, unless stated otherwise.
Mammalian cell transfection
Transfection complexes were prepared by mixing DNA and PEI
(polyethylenimine, linear, 25 kD; DNA : PEI [w/w] = 1 : 2.5) in
Opti-MEM at room temperature for 20 min. For every 35 mm
culture dish, 10 μL PEI (1 μg μL
1
) was used to prepare a
500 μL transfection mixture. We used a pcDNA3 plasmid
harboring a mitochondria-localized mW asabi (Mito-mWasabi).
22
The prepared transfection mixture was then added to cells and
incubated for 2 h at 37 °C. Next, pre-warmed, fresh DMEM
containing 10% FBS was used to replace the transfection
medium. On the following day, cells were stained with NIMAP
before imaging.
Fluorescence imaging
Fluorescence microscopy was performed with a Leica SP5
inverted confocal fluorescence microscope with the spectral
imaging capability (Leica, Boston, MA), unless otherwise
stated. The instrument was equipped with a 488 nm laser
(1.5 mW at full power), a 514 nm laser (2 mW at full power),
and a 633 nm laser (1 mW at full power). A 40× water lens was
used for all imaging studies. Unless otherwise noted, NIMAP
was imaged using a 633 nm laser line at 15% laser power for
excitation, and the gain was set to 800 mV. The emission was
set from 650 nm to 800 nm. To record the emission spectrum
for NIMAP loaded into live-cell mitochondria, the emission
bandwidth was set at 10 nm and images were taken from
650 nm to 790 nm. To image mWasabi GFP, the excitation was
set at 488 nm with 10% laser power, and the gain was set to
900 mV. The emission channel was set from 495 nm to
550 nm. To image Rhodamine 123, the excitation was set at
514 nm with 15% laser power, and the gain was set to 900 mV.
The emission was set from 525 nm to 570 nm. Time-lapse
images were acquired every minute. After the initial few
frames, oligomycin (MP Biomedicals, Solon, OH) was added to
a final concentration of 1 μgmL
1
. Several additional consecu-
tive images were acquired before adding FCCP (Enzo Life
Sciences, Farmingdale, NY) at a final concentration of 100 nM.
Additional frames were recorded. Similarly, we added H
2
O
2
(0.2%) to fresh HEK 293T cells loaded with NIMAP. Images
Paper Analyst
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were captured at 1 min intervals to observe the changes in
mitochondrial membrane potential and morphology. Cell
images were processed with Leica Application Suite (LAS) or
ImageJ.
Mouse imaging
Human prostate cancer PC3 cells were cultured as previously
described.
23
At 100% confluency, 4 × 10
6
cells per group were
stained with NIMAP (10 μM) or Rhodamine 123 (10 μM) for 1 h
at 37 °C. After staining with NIMAP, cells were washed once
with 5 ml of PBS. For collection, cells were suspended in the
culture medium and centrifuged at 3000 rpm for 2 min. Cells
were then resuspended in PBS and centrifuged at 3000 rpm for
2 min. The staining of cells with Rhodamine 123 was similar,
except two additional washes were carried out prior to the col-
lection of cells in order to remove excess Rhodamine 123.
Briefly, cells were allowed to incubate with the fresh culture
medium at 37 °C for 1 h, and this process was repeated before
collection of cells. Next, 4 × 10
6
cells were resuspended in
100 μL PBS and injected subcutaneously into the back of
immunocompromised NOD scid gamma mice (Jackson Labora-
tory, Stock # 005557) after air removal. Mice were imaged
several times over a week, using a Luminescence Dark Box
equipped with light-emitting diodes (LED) for excitation. To
record the fluorescence of NIMAP, we used a 630 nm LED
array, a 650/40 nm bandpass excitation filter, and a 695/30 nm
bandpass emission filter. To record the fluorescence of Rhoda-
mine 123, we used a 490 nm LED array, a 500/40 nm bandpass
excitation filter, and a 550/30 nm bandpass emission filter.
Images were processed with ImageJ.
Results and discussion
Designing and preparation of NIMAP
Our development of the new NIR MMP probes originated from
a Förster resonance energy transfer (FRET) experiment in our
laboratory, which used a QSY-21 core structure as a dark
quencher.
24
QSY-21 (1 in Fig. 1) is essentially nonfluorescent
(zero quantum yield) and has a high extinction coecient
(90 000 M
1
cm
1
at 661 nm).
25
It has been widely utilized as
an ecient nonfluorescent FRET acceptor for far-red and NIR
fluorophores. We prepared a QSY-21 labeled biotin derivative
(2), and observed mitochondria-like organellar fluorescence in
cultured mammalian cells stained with 100 μM of the dye (ESI
Fig. S1A). In contrast, incubating HEK 293T cells with a
neutral zwitterionic sulfonic acid form of QSY-21 (3) did not
induce significant NIR mitochondrial fluorescence (Fig. S1B).
Unstained HEK 293T cells also had no autofluorescence at this
spectral region (Fig. S1C and S1D). Considering the facts that
QSY-21 labeled biotin has some degrees of lipophilicity and a
positive charge, we reasoned that it was partially sequestered
into the lipid bilayers of cells, followed by migration into mito-
chondria and fluorescence activation in this hydrophobic,
viscous microenvironment.
In light of the potential use of QSY-21 derivatives as mito-
chondrial dyes, we designed a new molecule (4) containing a
hexanoamide connected to the QSY-21 core through a pipera-
zine sulfonamide linker. A short C6 fatty acid chain was
chosen to enhance the lipophilicity while retaining adequate
solubility of the resultant molecule in neutral aqueous solu-
tions for dye loading. This molecule was designated NIMAP
for its NIR, mitochondria-activatable property. A synthetic
route was developed to prepare NIMAP in 7 steps with 18%
overall yield from commercially available, inexpensive starting
materials (ESI Scheme S1).
Optimization of NIMAP loading into mammalian cells
We next loaded various concentrations of NIMAP into HEK
293T cells. No obvious cell death was observed upon 24 h
incubation with up to 100 μM NIMAP. We also studied the
impacts of dye concentrations and loading time on the
accumulation of NIMAP in mitochondria at 37 °C. To keep the
loading period as short as 45 min, 15 μM NIMAP in the cell
culture medium was needed to induce strong mitochondrial
fluorescence detectable by confocal microscopy with reason-
able instrumental settings (Fig. S2A). In comparison, 100 μM
2 was needed to gain similar images (Fig. S1A). These results
indicate that the fusion of the QSY-21 core with a C6 fatty acid
chain increases its partition into the lipid phase and its
accumulation in mammalian mitochondria.
The length of the loading period also aected the locali-
zation of NIMAP. With 1 μ M NIMAP, the mitochondrial fluo-
rescence continuously increased in a 2 h test window
(Fig. S2B), indicating that an equilibrium state for dye distri-
bution was not reached under these conditions. At 2 h, NIMAP
started to saturate mitochondria to also stain cytoplasmic
membrane (Fig. S2C). Due to the common practice favoring
short loading time, we used 1 μM NIMAP and a 45 min dye
incubation time in most of our following cell studies. No wash
step was needed to remove excess NIMAP in these experiments
because NIMAP was nonfluorescent in aqueous solutions. By
comparison, the background fluorescence of HEK 293T cells
stained with a conventional MMP dye, Rhodamine 123, was
strong before washing o
excess dye molecules (Fig. S3). We
also noticed a drastic dierence of NIMAP and Rhodamine
123 in terms of their photostability (Fig. 2). Almost no photo-
Fig. 1 Chemical structures of the dark quencher QSY-21 (1), a QSY-21
labeled biotin (2), a sulfonic acid form of QSY-21 (3), and NIMAP (4).
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bleaching was observed for NIMAP under the highest laser
power (100% of 1 mW at 633 nm), whereas Rhodamine 123-
stained cells photobleached quickly under a similar condition
(50% of 2 mW at 514 nm). The excellent photostability of
NIMAP makes it suitable for long-term time-lapse imaging.
Mitochondrial localization of NIMAP
To further confirm the mitochondrial localization of NIMAP,
we loaded the molecule into HEK 293T cells simultaneously
expressing a green fluorescent mWasabi protein fused to a
mitochondrial localization tag (Mito-mWasabi).
22
Dual colour
imaging verified the colocalization of mWasabi and NIMAP in
the mitochondria (Fig. 3). The ability to use NIMAP and a
green fluorescent probe for dual color imaging is important
because many existing, popular fluorescent sensors are based
on GFP, whereas the fluorescence excitation or emission of
common MMP probes, such as Rhodamine 123 and JC-1,
11
is
partially overlapped with that of GFP.
Fluorescence activation of NIMAP by biolipids
To determine whether the observed mitochondrial fluo-
rescence was due to the activation of NIMAP by the lipid mem-
brane, we prepared NIMAP-containing liposomes, following a
previously reported liposome-synthetic procedure.
21
The fluo-
rescence of NIMAP-containing liposomes was high (Fig. 4).
The excitation maximum is 650 nm and the emission
maximum is 710 nm. In comparison, NIMAP alone was essen-
tially non-fluorescent in aqueous solutions. Moreover, the
emission profile of NIMAP in liposomes was similar to that of
NIMAP in live-cell mitochondria (Fig. S4A). A previous mole-
cular dynamics (MD) study suggests that QSY-21 fluorescence
is quenched by ring rotations and electron transfer in the
excited state.
26
It is possible that the packed, viscous lipid
microenvironment may restrict the rotations and electron
transfer of NIMAP to enhance its fluorescence. We also
measured the absorbance of NIMAP in an aqueous solution,
which was similar to the profile of NIMAP fluorescence exci-
tation in liposomes (Fig. 4). Moreover, the absorbance was
insensitive to pH changes from pH 4 to 10 (Fig. S4B). We
further characterized the in vitro fluorescence of NIMAP in a
series of aqueous buers containing 90% (v/v) glycerol, and
the fluorescence intensities were essentially unchanged from
pH 4 to 10. These results are aligned with the conventional use
of QSY-21 as a dark quencher and our new finding that the
fluorescence of QSY-21 is greatly enhanced upon being inte-
grated into the mitochondrial membrane.
Fig. 2 Photostability of NIMAP (red) and rhodamine 123 (yellow) in HEK
293T cells under a confocal microscope.
Fig. 3 Colocalization of mitochondrial mWasabi (green channel) and
NIMAP (NIR channel) in HEK 293T cells (scale bar: 30 μm).
Fig. 4 Fluorescence excitation (open circle) and emission (closed
circle) spectra of NIMAP in liposomes (red) or an aqueous solution
(blue). The normalized absorbance of NIMAP in the aqueous solution is
also shown (gray line).
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Journal ArticleDOI
TL;DR: This critical review may promote the development of more robust MMP fluorescent probes, providing more powerful tools for researching basic biology to improve cancer diagnoses and treatments in clinics in the future.
Abstract: Mitochondrial membrane potential (MMP), as a key indicator of mitochondrial function, can reflect the cellular health status. The dysfunction of MMP, even the subtle unusual changes, can greatly affect the intracellular bioactivities, causing various diseases, such as keratitis, diabetes, Alzheimer’s disease and even cancer. Thus, detecting the variation of MMP has great significance for biological research and medical diagnosis. Due to the advantages of real-time and in-situ monitoring, a variety of organic fluorescent probes have been developed in recent years for the detection of MMP. However, this interesting and frontier topic has not been reviewed so far. In this review, we will focus on several kinds of recent organic fluorescent probes that have provided insights into MMP detection, including design concepts, responding mechanisms and applications of the representative examples. In addition, there are still some shortcomings and limitations of existing fluorescent probes on MMP detection, such as susceptible to interference, unquantifiable detection, and lack of clinical application. This critical review may promote the development of more robust MMP fluorescent probes, providing more powerful tools for researching basic biology to improve cancer diagnoses and treatments in clinics in the future.

13 citations


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Journal ArticleDOI
TL;DR: The description outlined here facilitates the understanding of factors that favour mitochondrial ROS production and develops better methods to measure mitochondrial O2•− and H2O2 formation in vivo, as uncertainty about these values hampers studies on the role of mitochondrial ROS in pathological oxidative damage and redox signalling.
Abstract: The production of ROS (reactive oxygen species) by mammalian mitochondria is important because it underlies oxidative damage in many pathologies and contributes to retrograde redox signalling from the organelle to the cytosol and nucleus. Superoxide (O2•−) is the proximal mitochondrial ROS, and in the present review I outline the principles that govern O2•− production within the matrix of mammalian mitochondria. The flux of O2•− is related to the concentration of potential electron donors, the local concentration of O2 and the second-order rate constants for the reactions between them. Two modes of operation by isolated mitochondria result in significant O2•− production, predominantly from complex I: (i) when the mitochondria are not making ATP and consequently have a high Δp (protonmotive force) and a reduced CoQ (coenzyme Q) pool; and (ii) when there is a high NADH/NAD+ ratio in the mitochondrial matrix. For mitochondria that are actively making ATP, and consequently have a lower Δp and NADH/NAD+ ratio, the extent of O2•− production is far lower. The generation of O2•− within the mitochondrial matrix depends critically on Δp, the NADH/NAD+ and CoQH2/CoQ ratios and the local O2 concentration, which are all highly variable and difficult to measure in vivo. Consequently, it is not possible to estimate O2•− generation by mitochondria in vivo from O2•−-production rates by isolated mitochondria, and such extrapolations in the literature are misleading. Even so, the description outlined here facilitates the understanding of factors that favour mitochondrial ROS production. There is a clear need to develop better methods to measure mitochondrial O2•− and H2O2 formation in vivo, as uncertainty about these values hampers studies on the role of mitochondrial ROS in pathological oxidative damage and redox signalling.

5,363 citations


Journal ArticleDOI
TL;DR: Measurement of both respiration and potential during appropriate titrations enables the identification of the primary sites of effectors and the distribution of control, allowing deeper quantitative analyses, as discussed in the present review.
Abstract: Assessing mitochondrial dysfunction requires definition of the dysfunction to be investigated. Usually, it is the ability of the mitochondria to make ATP appropriately in response to energy demands. Where other functions are of interest, tailored solutions are required. Dysfunction can be assessed in isolated mitochondria, in cells or in vivo, with different balances between precise experimental control and physiological relevance. There are many methods to measure mitochondrial function and dysfunction in these systems. Generally, measurements of fluxes give more information about the ability to make ATP than do measurements of intermediates and potentials. For isolated mitochondria, the best assay is mitochondrial respiratory control: the increase in respiration rate in response to ADP. For intact cells, the best assay is the equivalent measurement of cell respiratory control, which reports the rate of ATP production, the proton leak rate, the coupling efficiency, the maximum respiratory rate, the respiratory control ratio and the spare respiratory capacity. Measurements of membrane potential provide useful additional information. Measurement of both respiration and potential during appropriate titrations enables the identification of the primary sites of effectors and the distribution of control, allowing deeper quantitative analyses. Many other measurements in current use can be more problematic, as discussed in the present review.

1,727 citations


Journal ArticleDOI
TL;DR: This paper summarizes exclusively scalable techniques and focuses on strengths and limitations in respect to industrial applicability and regulatory requirements concerning liposomal drug formulations based on FDA and EMEA documents.
Abstract: Liposomes, sphere-shaped vesicles consisting of one or more phospholipid bilayers, were first described in the mid-60s. Today, they are a very useful reproduction, reagent, and tool in various scientific disciplines, including mathematics and theoretical physics, biophysics, chemistry, colloid science, biochemistry, and biology. Since then, liposomes have made their way to the market. Among several talented new drug delivery systems, liposomes characterize an advanced technology to deliver active molecules to the site of action, and at present, several formulations are in clinical use. Research on liposome technology has progressed from conventional vesicles to ‘second-generation liposomes’, in which long-circulating liposomes are obtained by modulating the lipid composition, size, and charge of the vesicle. Liposomes with modified surfaces have also been developed using several molecules, such as glycolipids or sialic acid. This paper summarizes exclusively scalable techniques and focuses on strengths, respectively, limitations in respect to industrial applicability and regulatory requirements concerning liposomal drug formulations based on FDA and EMEA documents.

1,710 citations


Journal ArticleDOI
TL;DR: Accumulation of Ca2+ into mitochondria regulates mitochondrial metabolism and causes a transient depolarisation of mitochondrial membrane potential, and alteration of spatiotemporal characteristics of cellular [Ca2+]c signalling and downregulates mitochondrial metabolism.
Abstract: While a pathway for Ca2+ accumulation into mitochondria has long been established, its functional significance is only now becoming clear in relation to cell physiology and pathophysiology. The observation that mitochondria take up Ca2+ during physiological Ca2+ signalling in a variety of cell types leads to four questions: (i) ‘What is the impact of mitochondrial Ca2+ uptake on mitochondrial function?’ (ii) ‘What is the impact of mitochondrial Ca2+ uptake on Ca2+ signalling?’ (iii) ‘What are the consequences of impaired mitochondrial Ca2+ uptake for cell function?’ and finally (iv) ‘What are the consequences of pathological [Ca2+]c signalling for mitochondrial function?’ These will be addressed in turn. Thus: (i) accumulation of Ca2+ into mitochondria regulates mitochondrial metabolism and causes a transient depolarisation of mitochondrial membrane potential. (ii) Mitochondria may act as a spatial Ca2+ buffer in many cells, regulating the local Ca2+ concentration in cellular microdomains. This process regulates processes dependent on local cytoplasmic Ca2+ concentration ([Ca2+]c), particularly the flux of Ca2+ through IP3-gated channels of the endoplasmic reticulum (ER) and the channels mediating capacitative Ca2+ influx through the plasma membrane. Consequently, mitochondrial Ca2+ uptake plays a substantial role in shaping [Ca2+]c signals in many cell types. (iii) Impaired mitochondrial Ca2+ uptake alters the spatiotemporal characteristics of cellular [Ca2+]c signalling and downregulates mitochondrial metabolism. (iv) Under pathological conditions of cellular [Ca2+]c overload, particularly in association with oxidative stress, mitochondrial Ca2+ uptake may trigger pathological states that lead to cell death. In the model of glutamate excitotoxicity, microdomains of [Ca2+]c are apparently central, as the pathway to cell death seems to require the local activation of neuronal nitric oxide synthase (nNOS), itself held by scaffolding proteins in close association with the NMDA receptor. Mitochondrial Ca2+ uptake in combination with NO production triggers the collapse of mitochondrial membrane potential, culminating in delayed cell death.

1,014 citations


Journal ArticleDOI
Lan Bo Chen1

949 citations