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ARTICLE
Received 11 Feb 2014 | Accepted 23 Apr 2014 | Published 29 May 2014
Separating NADH and NADPH fluorescence
in live cells and tissues using FLIM
Thomas S. Blacker
1,2,3
, Zoe F. Mann
2,4
, Jonathan E. Gale
2,4
, Mathias Ziegler
5
, Angus J. Bain
3
,
Gyorgy Szabadkai
2,6,
* & Michael R. Duchen
2,
*
NAD is a key determinant of cellular energy metabolism. In contrast, its phosphorylated form,
NADP, plays a central role in biosynthetic pathways and antioxidant defence. The reduced
forms of both pyridine nucleotides are fluorescent in living cells but they cannot be
distinguished, as they are spectrally identical. Here, using genetic and pharmacological
approaches to perturb NAD(P)H metabolism, we find that fluorescence lifetime imaging
(FLIM) differentiates quantitatively between the two cofactors. Systematic manipulations to
change the balance between oxidative and glycolytic metabolism suggest that these states do
not directly impact NAD(P)H fluorescence decay rates. The lifetime changes observed in
cancers thus likely reflect shifts in the NADPH/NADH balance. Using a mathematical model,
we use these experimental data to quantify the relative levels of NADH and NADPH in
different cell types of a complex tissue, the mammalian cochlea. This reveals NADPH-enri-
ched populations of cells, raising questions about their distinct metabolic roles.
DOI: 10.1038/ncomms4936
OPEN
1
Centre for Mathematics and Physics in the Life Sciences and Experimental Biology, University College London, London WC1E 6BT, UK.
2
Research
Department of Cell & Developmental Biology, University College London, London WC1E 6BT, UK.
3
Department of Physics and Astronomy, University College
London, London WC1E 6BT, UK.
4
UCL Ear Institute, University College London, London WC1X 8EE, UK.
5
Department of Molecular Biology, University of
Bergen, N-5008 Bergen, Norway.
6
Department of Biomedical Sciences, University of Padua and CNR Neuroscience Institute, Padua 35121, Italy. * These
authors contributed equally to this work. Correspondence and requests for materials should be addressed to M.R.D. (email: m.duchen@ucl.ac.uk).
NATURE COMMUNICATIONS | 5:3936 | DOI: 10.1038/ncomms4936 | www.nature.com/naturecommunications 1
& 2014 Macmillan Publishers Limited. All rights reserved.
T
he cellular redox state is the central regulator of energy
production and intermediary metabolism, playing a
crucial role in health and disease
1
. The nicotinamide
adenine dinucleotide (NAD
þ
/NADH) and nicotinamide adenine
dinucleotide phosphate (NADP
þ
/NADPH) redox couples are
the major determinants of redox state in the cell. However, these
engage in distinct metabolic pathways. NAD drives ATP
production in the cytosol by glycolysis and in the mitochondria
by oxidative phosphorylation, while the phosphorylated analogue
NADP governs lipid, amino acid and nucleotide biosynthetic
pathways and the defence against reactive oxygen species by
glutathione (GSH)
2
. Free radical generation is therefore
determined by the redox state of NAD, while NADP redox
state is key to antioxidant defence
3
. The relative abundance of the
two pyridine nucleotides and their redox balance thus mediates
cell fate in a wide range of diseases, including cancer, diabetes and
neurodegeneration. Quantifying their behaviour is therefore
essential in understanding the role of metabolism in these
diseases. However, separating the contributions of the two pools
in intact tissues has proven technically challenging
4
.
In the 1960s, Britton Chance et al.
5
showed that live tissues
illuminated with ultraviolet light emit blue fluorescence, arising
primarily from mitochondrial NADH. The nicotinamide moiety
of NADH absorbs light of wavelength 340
±
30 nm and emits
fluorescence at 460
±
50 nm. As NADP is phosphorylated at a
remote site of the molecule, the fluorescence properties of the
nicotinamide ring of NADPH are identical to those of NADH
6,7
.
Thus, changes in autofluorescence intensity may reflect changes
in either [NADH] or [NADPH], often denoted as NAD(P)H to
indicate the uncertain origin of the signal
8,9
.
Fluorescence lifetime imaging (FLIM) allows the study
of NAD(P)H photochemistry inside living tissue
10–12
.
This technique measures the rates of fluorescence decay of
NAD(P)H, an excited-state process occurring over nanosecond
time scales, which is highly sensitive to the immediate
environment of the fluorophore. We have therefore investigated
whether the fluorescence lifetime of cellular NADPH differs from
that of cellular NADH, reflecting the different set of enzymes to
which the two cofactors bind. Indeed, a substantial literature
documents variations in the fluorescence lifetime of NAD(P)H
in a range of physiological and pathological conditions
13–18
.
However, despite the potential clinical applications of
autofluorescence lifetime measurements, such as delineating the
boundaries of accessible cancers
14
, the biochemical basis for these
variations remains unknown. Moreover, the issue of whether
FLIM may permit separation of NADH and NADPH
fluorescence to resolve cellular specializations and dynamics
in intact tissues is yet to be addressed. In the present study,
we have explored the fluorescence decay properties of NADH and
NADPH in living cells and tissues. Combined with computational
and mathematical modelling, we have found that NAD(P)H
fluorescence lifetime characteristics discriminate between NADH
and NADPH. This provides a unique approach to identify cells
within complex tissues that are enriched in NADPH, thus raising
questions about their metabolic roles and specialization. We have
also analysed the impact of altered metabolic state on NAD(P)H
fluorescence decay characteristics, helping to place changes in
lifetimes observed in transformed neoplastic cells on a firm
biochemical footing.
Results
NAD(P)H fluorescence decay reflects bound NADPH/NADH
ratio. We have previously shown that the fluorescence lifetime
of NADPH is identical to that of NADH in solution
19
,
demonstrating that fluorescence from the free, unbound
pyridine nucleotides cannot be discriminated on the basis of
their fluorescence lifetime. However, it has long been known that
this value increases from 0.3–0.8 to 1–6.5 ns on binding to an
enzyme, depending on the target to which the cofactor binds
10,20
,
as well as by the simultaneous presence of substrate molecules
21
.
Canonical NAD(P)H FLIM studies in live cells and tissues resolve
two fluorescence lifetimes at each pixel; one of the order of 0.4 ns
and the other larger and more variable at B2 ns or more
11,12,22
.
These represent the freely diffusing (t
free
) and enzyme-bound
NAD(P)H (t
bound
) pools, respectively, confirmed by time-
resolved anisotropy imaging studies
23
. As the distribution of
bound NAD(P)H species will inevitably be more heterogeneous
than implied by the single long lifetime, we suggest that t
bound
is a
weighted mean of the fluorescence lifetimes of enzyme-bound
species, a conclusion supported by computational modelling (see
Supplementary Fig. 1a–m and Supplementary Note 1). It is
primarily this parameter that has previously been observed to
vary with changes in metabolism
11,24–26
.
Next, to understand the effect of varying [NADPH]/[NADH]
ratios on t
bound
, we acquired FLIM images of NAD(P)H in live
HEK293 cells in which NADPH levels were genetically and
pharmacologically manipulated. In wild-type cells, the fluores-
cence decays obtained at each pixel required a two-component
model to yield good fits (Fig. 1a,b), as expected, with w
2
r
values
close to unity. As such, addition of further components was not
appropriate, to avoid inaccurate weightings due to overfitting
27
.
Identical fluorescence decay parameters were observed in the
cytosol and mitochondria, with mean (
±
s.d.) values of
t
bound
¼ 2.7
±
0.2 ns, a
bound
¼ 0.19
±
0.01 (fraction of bound
NAD(P)H) and t
free
¼ 0.36
±
0.04 ns (Supplementary Table 1).
Interestingly, t
bound
was significantly smaller in the nucleus
at 2.3
±
0.2 ns (P ¼ 1E 8, two-tailed Student’s t-test, n ¼ 17
images).
To explain reported variations in t
bound
, we hypothesized that,
since NADH and NADPH are associated with different binding
site structures inside the cell
28
, the measured t
bound
value could
reflect the proportion of the two cofactors present. We therefore
measured NAD(P)H fluorescence lifetimes in HEK293 cell lines
in which NAD
þ
kinase (NADK) was either overexpressed
(NADK þ ) or knocked down (NADK )
29
. NADK is the key
determinant of NADPH concentration inside the cell, and its
overexpression results in a 10- to 15-fold higher [NADPH] in
NADK þ relative to NADK cells, leaving [NADH] relatively
unaffected
29
. This was reflected by an NAD(P)H fluorescence
intensity B10-fold brighter in NADK þ compared with
NADK cells (Supplementary Fig. 2b). FLIM revealed that
t
bound
was significantly larger in the NADK þ cells
(t
bound
¼ 3.6
±
0.2 ns in the mitochondria, 3.8
±
0.2 ns in the
cytosol) compared with the NADK cells (t
bound
¼ 2.7
±
0.1 ns
in both subcellular regions, P ¼ 1E 11 and P ¼ 5E 12,
two-tailed Student’s t-test, n ¼ 9, Fig. 1c and Supplementary
Table 1).
These data suggest that increased concentrations of bound
NADPH result in increased values of t
bound
. To verify this
interpretation, we explored the impact of epigallocatechin gallate
(EGCG) on the NAD(P)H FLIM parameters. EGCG is a potent
competitive inhibitor of NADPH binding
30
with no effect
on NADH binding indicated by the BRENDA database
31
.
Preferential competition for NADPH-binding sites by EGCG
would decrease the bound NADPH population, leaving the
bound NADH population unaffected. The value of a
bound
should
therefore be more sensitive to EGCG treatment in NADK þ cells
than in NADK cells, as bound NADPH will form a greater
fraction of the total population of enzyme-bound NAD(P)H
species. As predicted, a
bound
decreased significantly from
0.18
±
0.02 to 0.13
±
0.02 in both the mitochondria and cytosol
ARTICLE NATURE COMMUNICATIONS | DOI: 10.1038/ncomms4936
2 NATURE COMMUNICATIONS | 5:3936 | DOI: 10.1038/ncomms4936 | www.nature.com/naturecommunications
& 2014 Macmillan Publishers Limited. All rights reserved.
(P ¼ 4E 10, two-tailed Student’s t-test, n ¼ 9) on EGCG
treatment in NADK þ cells but remained constant in NADK
cells, confirming the NADPH specificity of this compound (see
Fig. 1d). Exposure to EGCG decreased t
bound
to 3.1
±
0.2 ns
in both the mitochondria and cytosol of NADK þ cells
(P ¼ 2E 12, two-tailed Student’s t-test, n ¼ 9) and to
2.5
±
0.2 ns in their nuclei (P ¼ 3E 7, two-tailed Student’s
t-test, n ¼ 9), and did not significantly affect NADK cells
(Fig. 1c). In addition, treatment with this compound did not
affect t
free
(Supplementary Fig. 2c,d). Thus, these data are
consistent with a model where t
bound
reports the amount of
enzyme-bound NADPH relative to bound NADH.
These data suggest that t
bound
can be used to quantify enzyme-
bound NADPH/NADH ratios. We therefore developed a
numerical model to quantify this phenomenon and generate
predictions (see Supplementary Fig. 1k–o and Supplementary
Note 2). By combining the enzyme-bound NAD(P)H fluores-
cence lifetimes measured in the NADK þ and NADK cells, the
biochemically quantified [NADH] and [NADPH] values in each
cell line
29
and a mathematical model in which NADH and
NADPH were assumed to possess discrete and distinct
fluorescence lifetimes when bound inside the cell, we found
that t
bound
would describe the [NADPH]/[NADH] ratio by
NADPH½
NADH½
¼
t
bound
nsðÞ1:5
4:4 t
bound
nsðÞ
ð1Þ
Application of this model to the NADK þ and NADK data
(see Supplementary Note 3) showed that the concentration of
bound NADH remained constant on EGCG application, while
the concentration of bound NADPH decreased B3-fold with
small differences in each subcellular compartment, supporting
our hypothesis.
Lifetime changes reflect mechanism of metabolic perturbation.
In previous work
11
, shortening of t
bound
observed in tumours has
been attributed to a shift from oxidative to glycolytic metabolism
which occurs in many cancers, the so-called Warburg effect. To
investigate this hypothesis in the light of the results described
here, FLIM images of wild-type HEK293 cells were acquired
following a range of manipulations that alter the balance of ATP
production by aerobic or anaerobic pathways. Dependence on
glycolysis was achieved by inhibition of mitochondrial oxidative
phosphorylation using rotenone (10 mM) or uncoupling using
Mitochondria
NADK+
NADK+
*
*
*
*
*
*
*
*
*
*
*
*
NADK+ NADK+
w/EGCG
NADK–
w/EGCG
NADK–
NADK–
w/EGCG
NADK+ NADK+
w/EGCG
NADK–
NADK–
NADK–
ControlControl
w/EGCGw/EGCG
4.0
2.0
0.3
bound
(ns)
4.0
3.5
3.0
2.5
2.0
0.0
0.0
0.0
0.3
0.2
0.1
Cytosol Nucleus
bound
IRF
Biexponential fit
IRF
Monoexponential fit
Time (ns)
500
0
10
–10
500
0
0
10
–10
0
0
123
4567
Weighted
residual
χ
2
= 1.1 χ
2
= 5.0
Time (ns)
0
123
456
7
Photon counts
Photon counts
Figure 1 | s
bound
reflects the enzyme-bound NADPH/NADH ratio in intact cells. (a,b) A biexponential decay model adequately described the NAD(P)H
fluorescence decay measured in wtHEK293 cells (IRF, instrument response function). The mean w
2
r
was 1.24
±
0.08 compared with 4
±
1 with a
monoexponential fit (representative data from n ¼ 17 experiments). (c,d) Representative colour-coded images and mean t
bound
and a
bound
values in
NADK þ and NADK HEK293 cells prior and following treatment with EGCG (100 mM), a competitive inhibitor of NADPH binding. Scale bar, 20 mm.
Error bars indicate
±
s.d., *Po0.05 (two-tailed Student’s t-test, n ¼ 9).
NATURE COMMUNICATIONS | DOI: 10.1038/ncomms4936 ARTICLE
NATURE COMMUNICATIONS | 5:3936 | DOI: 10.1038/ncomms4936 | www.nature.com/naturecommunications 3
& 2014 Macmillan Publishers Limited. All rights reserved.
FCCP (1 mM). The cells were driven to a more oxidative
phenotype by inhibition of glycolysis by glucose deprivation in
the presence of deoxyglucose (10 mM), while pyruvate (1 mM)
or lactate (10 mM) were provided as mitochondrial-specific
substrates.
Ultraviolet confocal microscopy was used to establish the time
taken for the redox state of the NAD(P) pool to reach a new
steady state following each treatment (Fig. 2a–d). Inhibition of
glycolysis decreased the cellular NAD(P)H fluorescence intensity
by 26
±
6 and 22
±
8% with pyruvate and lactate supplied,
respectively. Rotenone increased steady-state NAD(P)H fluores-
cence intensity by 20
±
5% while FCCP caused a decrease by
38
±
2% (each n ¼ 3). FLIM images were acquired at the steady-
state fluorescence intensity levels following each treatment
(Fig. 2e–g and Supplementary Table 2). Rotenone caused t
bound
to decrease significantly from 2.7
±
0.2 to 2.52
±
0.05 ns in both
the mitochondria and cytosol (P ¼ 6E 4, two-tailed Student’s
t-test, n ¼ 9), suggesting that this treatment caused the concen-
tration of NADH present in the cell to increase relative to the
concentration of NADPH, following inhibition of NADH
oxidation by complex I. In contrast, t
bound
did not change in
response to FCCP (P40.05, two-tailed Student’s t-test, n ¼ 13).
This lack of change in t
bound
suggested that uncoupling caused
the oxidation of both NAD and NADP pools in equal measure.
Increased oxidation of the NAD pool was to be expected on
uncoupling due to the increased complex I activity, and the equal
oxidation of the NADP pool was likely caused by the action of the
mitochondrial transhydrogenase. In respiring mitochondria, this
inner mitochondrial membrane protein transfers hydride from
NADH to NADP
þ
powered by translocation of protons from the
intermembrane space to the mitochondrial matrix
32
. However, on
uncoupling, NADPH is oxidized and the hydride is passed to
Time (min)
Time (min)
NAD(P)H intensity (a.u.)
150
50
0
100
NAD(P)H intensity (a.u.)
150
50
0
100
NAD(P)H intensity (a.u.)
150
50
0
100
NAD(P)H intensity (a.u.)
150
50
0
100
Time (min)
Time (min)
Glucose Rotenone
FCCP Pyruvate Lactate
(Glucose deprivation)
4.0
2.0
bound
(ns)
bound
(ns)
bound
bound
0.3
0.0
Mitochondria Cytosol Nucleus
*
0.2
0.3
0.1
0.0
*
*
*
*
Glucose Rotenone FCCP Pyruvate Lactate
(Glucose deprivation)
Glucose Rotenone FCCP Pyruvate Lactate
(Glucose deprivation)
*
*
*
*
*
*
*
*
*
Rotenone
Pyruvate
0246810
02468
10
0246810
0246810
Lactate
FCCP
0.0
2.0
2.5
3.0
4.0
3.5
Figure 2 | NAD(P)H fluorescence decay responses to metabolic perturbation are mechanism dependent. (a–d) Time series of NAD(P)H
fluorescence intensity following treatments chosen to perturb oxidative (a: respiratory chain inhibition by rotenone, b: uncoupling with FCCP) and glycolytic
metabolism (glucose replaced by deoxyglucose with c: pyruvate or d: lactate supplied as substrate). (e) Colour-coded images and (f,g) quantification
of changes in t
bound
and a
bound
on application of treatment. Scale bar, 20 mm. Error bars indicate
±
s.d., *Po0.05 (two-tailed Student’s t-test, n ¼ 9).
ARTICLE NATURE COMMUNICATIONS | DOI: 10.1038/ncomms4936
4 NATURE COMMUNICATIONS | 5:3936 | DOI: 10.1038/ncomms4936 | www.nature.com/naturecommunications
& 2014 Macmillan Publishers Limited. All rights reserved.
NAD
þ
to produce NADH
33
. Relative to controls, glucose
deprivation in the presence of either pyruvate or lactate supply
failed to significantly alter t
bound
(P40.05, two-tailed Student’s
t-test, both n ¼ 9). However, the mean value of t
bound
in both the
mitochondria and cytosol of 2.70
±
0.01 ns in the presence of
pyruvate was significantly larger than its value of 2.57
±
0.04 ns in
the presence of lactate (P ¼ 7E 6, two-tailed Student’s t-test,
n ¼ 9). As lactate dehydrogenase promotes NADH production
during the conversion of lactate to pyruvate, the significantly
smaller t
bound
value in the presence of lactate relative to pyruvate
supports the interpretation developed in this work that this
parameter reflects NADPH/NADH ratio, assuming NADPH
production was identical in the presence of the two substrates.
While changes in t
bound
, and thus the NADPH/NADH ratio,
reflected the specific treatment causing a defect in OXPHOS
or glycolysis, we observed that each treatment causing a net
oxidation of the combined NAD(P) pools caused a
bound
to
increase (Fig. 2g). This is in support of previous suggestions that
this parameter, reflecting the enzyme-bound population fraction
of NAD(P)H, reports acute changes in the redox state of the
cell
25
. However, confirming whether this parameter reflects
phenotypic differences in the redox state of different cell types
requires further investigation. Interestingly, each oxidizing
treatment also caused an increase in t
free
(Supplementary
Fig. 2e,f). However, as these lifetimes lie close to the time
resolution of the FLIM system, such small differences may be an
artefact of the fitting process, such as an interdependency between
t
free
and a
bound
(Supplementary Fig. 1j).
The lack of significant change in t
bound
in response to FCCP
treatment or replacement of glucose with pyruvate or lactate,
along with the very small, if significant, change in this parameter
following rotenone treatment relative to the pathophysiological
variations reported in the literature, suggests that changes in
NAD(P)H fluorescence decay cannot be simply attributed to
alterations in the balance between oxidative and glycolytic
metabolism. Accordingly, no correlation between t
bound
and the
balance of ATP production by glycolytic or oxidative means was
observed by measuring the rates of oxygen consumption and
lactate release in the wild-type, NADK þ and NADK HEK293
cell lines (see Supplementary Fig. 3 and Supplementary Tables 3
and 4). Altogether, these data strongly suggest that any variation
in the NAD(P)H fluorescence decay parameter t
bound
will be
specific to the mechanism of metabolic perturbation, such as
mitochondrial dysfunction caused either by respiratory chain
inhibition or uncoupling, or the utilization of different substrates
following inhibition of glycolysis. Such specificity has the
potential to aid the study of the variety of possible metabolic
rearrangements that may occur in cancer and other pathologies.
FLIM reveals metabolic variations in complex tissues. The
abundance of NADPH and NADH in live cells can be measured
Untreated
OHCs
Intensity Intensity
OHCs
OHCs
OHCs
OHCs
OHCs
OPCs
OPCs
OPCs
OPCs
OHCs
OPCs
OPC
OPCs
w/EGCG
w/EGCG
*
*
*
*
*
bound
(ns)
4.0
3.0
2.0
2.0 4.0 4.02.0
MCB fluorescence (a.u.)
0.0
0.5
1.0
1.5
2.0
*
bound
0.0
0.1
0.2
0.3
0.0
bound
(ns)
bound
(ns)
Figure 3 | Supporting cells in the mammalian cochlea exhibit increased enzyme-bound NADPH. (a) Mean t
bound
and (b) a
bound
values in outer hair
cells (OHC’s) and adjacent outer pillar ‘supporting’ cells (OPC’s) under control conditions and following application of EGCG (200 mM). Error bars
indicate
±
s.d., *Po0.05 (two-tailed Student’s t-test, n ¼ 11). (c,d) Corresponding representative FLIM images colour coded for the mean parameter
value in each cell. Scale bar, 25 mm. ( e) Schematic diagram showing organ of Corti in the cochlear explants, indicating the positions of OHC’s and OPC’s.
(f) Mean fluorescence intensity in OPC’s and OHC’s. Error bars indicate
±
s.d., *Po0.05 (Wilcoxon signed-rank test, n ¼ 17). (g) Representative image
following MCB staining for GSH concentration.
NATURE COMMUNICATIONS | DOI: 10.1038/ncomms4936 ARTICLE
NATURE COMMUNICATIONS | 5:3936 | DOI: 10.1038/ncomms4936 | www.nature.com/naturecommunications 5
& 2014 Macmillan Publishers Limited. All rights reserved.
in tissues by biochemical means such as high performance liquid
chromatography
29,34
. However, a microscopy-based imaging
approach permits measurement of NADH and NADPH in
discrete compartments within a cell or in different cell types
within a complex tissue. Indeed, we have observed consistently
smaller values of t
bound
in the nucleus compared with the rest of
the cell. Combining enzyme localization data
31
with the analysis
and model provided here, NAD(P)H FLIM indicates that nuclear
glucose metabolism favours NADH-producing glycolysis rather
than NADPH-producing pentose phosphate pathway (PPP),
resulting in a more oxidized NADP pool in this region (see
Supplementary Note 4).
To explore the application of NAD(P)H FLIM in multicellular
specimens, the redox metabolism of the mammalian cochlea was
investigated. The cochlea is a structure containing a highly
ordered system of functionally distinct cell types, which are easily
identifiable from their architecture (Fig. 3e). FLIM imaging of
cochlear explant cultures revealed that t
bound
differed markedly
between different cell types. The glia-like outer pillar ‘supporting’
cells (OPCs) exhibited an extended fluorescence lifetime of
3.5
±
0.1 ns, while the mean value of t
bound
measured in the
neighbouring sensory receptors, the outer hair cells (OHCs),
was significantly lower at 2.9
±
0.1 ns (P ¼ 2E 15, two-tailed
Student’s t-test, n ¼ 19, Fig. 3a–d). EGCG treatment decreased
t
bound
in supporting cells by 0.6
±
0.2 ns (P ¼ 1E 6, two-tailed
Student’s t-test, n ¼ 11) to a mean of 2.9
±
0.2 ns, but by only
0.2
±
0.2 ns (P ¼ 8E 4, two-tailed Student’s t-test, n ¼ 11) to a
mean of 2.7
±
0.1 ns in hair cells (Supplementary Table 5).
These data confirm that the differences in fluorescence lifetime
between cell types were a measure of differences in the bound
NADPH/NADH ratio.
Application of the mathematical model developed in this work
(Supplementary equations 10 and 11) implied that the total
concentration of reduced pyridine nucleotides were equal in the
OHCs and OPCs. In the OHCs, the absolute concentrations of
NADH and NADPH were equal at 1
±
0.2 a.u. However, in the
OPCs, the concentration of NADPH was significantly larger at
1.4
±
0.3 a.u. (P ¼ 4E 4, Wilcoxon signed-rank test, n ¼ 19),
with a correspondingly smaller concentration of NADH (0.7
±
0.2
a.u., P ¼ 2E 4, Wilcoxon signed-rank test, n ¼ 19). To assess the
functional significance of this finding, we measured the distribu-
tion of GSH in the tissue using monochlorobimane (MCB)
staining, since a major demand for NADPH in glial cells arises
from GSH turnover
35
. The glia-like OPCs showed significantly
higher [GSH] compared with OHCs (P ¼ 0.02, Wilcoxon signed-
rank test, n ¼ 17, Fig. 3f,g), consistent with a functional
requirement for NADPH enrichment. Thus, our interpretation
of t
bound
is consistent with alternative indicators of cellular redox
state.
Discussion
This work shows that FLIM can be used effectively to differentiate
between NADH and NADPH at the level of the single cell or
organelle. The results suggest that enzyme-bound NADPH
possesses a significantly larger fluorescence lifetime than
enzyme-bound NADH within the cellular environment, so that
the proportion of enzyme-bound NADPH and NADH present in
live tissue determines the lifetime of their combined fluorescence
decay. By making the simplifying assumption that bound NADH
and bound NADPH possess finite and distinct fluorescence
lifetimes inside the cell, the relative contribution of each cofactor
to the combined fluorescence signal could be calculated. With
excitation at 700 nm and a 435–485 nm detection window, the
intracellular fluorescence lifetimes of NADH and NADPH
were predicted to be 1.5
±
0.2 and 4.4
±
0.2 ns respectively. It is
reasonable to hypothesize that these conclusions can be extended
to all cell types as the fluorescence lifetime of NAD(P)H when
bound to an enzyme is determined by its local environment in the
binding site
21
, and the NADH and NADPH-binding sites are two
of the most highly conserved in all biology
36,37
. Indeed, we have
observed values of t
bound
that are similar in magnitude across a
range of cell types, including isolated ventricular cardiomyocytes
and neurons in culture or in brain slices (data not shown).
The analysis and model presented herein are consistent with
previously published NAD(P)H FLIM studies. In 2008, Niesner
et al.
38
performed a novel study on the decay of NAD(P)H
fluorescence in granulocytes in the presence of Aspergillus
fumigatus fungus. The parameter t
bound
was B2 ns within
the bulk cytosol of the granulocytes. However, localized
subplasmalemmal regions of the cytosol in contact with the
fungus displayed increased values of 3.7 ns. This was attributed to
a unique fluorescence lifetime of NADPH when bound to the
NADPH oxidase activated in response to pathogenic exposure.
However, the computational simulations performed here showed
that t
bound
is a weighted average of the fluorescence lifetimes of
the enzyme-bound NAD(P)H species present. For NADPH
oxidase alone to cause an increase in t
bound
as large as that
observed in the granulocytes would require this enzyme to be
present at a greater concentration than the NADH-binding
enzymes of the cytosol. The Model Organism Protein Expression
Database (MOPED)
39
within the GeneCards human gene
compendium
40
shows that expression of glyceraldehyde
3-phosphate dehydrogenase outweighs that of the NADPH-
binding subunit of NADPH oxidase (neutrophil cytosolic factor
2)
41
by around 100 to one in neutrophils. It is therefore more
likely that the NAD(P)H fluorescence lifetime observed in the
regions of the cytosol where NADPH oxidase was activated was
due to increased local NADPH production, as implied from the
results reported here. Indeed, the large quantities of superoxide
produced by NADPH oxidase requires plentiful supply of
NADPH. Activation of this enzyme is thus associated with
increased flux of glucose through the PPP
42
.
Changes in the fluorescence decay of NAD(P)H have recently
been observed in applications ranging from wound healing
17
to
stem cell differentiation
18
and necrotic deterioration of skin
15
to
staurosporine-induced apoptosis
16
. The large number of studies
reporting differences between the fluorescence lifetime of
NAD(P)H in healthy control cells and cells at different stages
of carcinogenesis
11,43,44
have prompted the design of clinical
instruments for the detection, diagnosis and staging of
accessible tumours using time-resolved autofluorescence
measurements
14,45–50
. While the Warburg effect is the most
well known of the metabolic shifts occurring during cancer
development, tumorigenesis is also associated with variations in
glucose flux through the PPP and various biosynthetic pathways
utilizing NADPH
51
. As none of the severe pharmacological
perturbations to cytosolic or mitochondrial ATP production
applied in this work could reproduce responses in t
bound
of the
magnitude caused by an increased NADPH/NADH ratio, our
results strongly suggest that NAD(P)H fluorescence lifetime
differences observed between healthy and pathological states
reflect shifts in the NADPH/NADH balance.
In this work, termination of oxidative phosphorylation by
uncoupling and of glycolysis by glucose deprivation could not be
resolved on the basis of the fluorescence decay parameters
measured under these conditions. In addition, two of the cell lines
studied here with similar fluorescence decay parameters were
shown to differ in their reliance on aerobic and anaerobic
metabolism (wild-type HEK293 and NADK ). This implies that
changes in the NAD(P)H fluorescence decay do not simply report
shifts between an oxidative or glycolytic phenotype, but reflect the
ARTICLE NATURE COMMUNICATIONS | DOI: 10.1038/ncomms4936
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& 2014 Macmillan Publishers Limited. All rights reserved.
differential response of the NAD and NADP pools following a
metabolic alteration. For example, both uncoupling and electron
transport chain inhibition terminate aerobic ATP production yet
have opposing effects on the overall NAD(P) redox state and
thus, perhaps on a
bound
. Changes in t
bound
will be induced by
metabolic transitions that cause divergent effects on NAD- and
NADP-associated pathways. Thus, neither parameter alone will
permit the detection of oxidative or glycolytic states using
NAD(P)H fluorescence decay measurements. However, a method
for assessing the pathways used for ATP production by
measuring autofluorescence may be possible by combining a
number of observables, including a
bound
and t
bound
, alongside
fluorescence intensity information and similar measurements
from flavoprotein fluorescence as part of a spectrally resolved
lifetime imaging tool
24
.
In the present study, we have investigated the canonical form
of FLIM in which fluorescence lifetimes t
free
and t
bound
, along
with the weighting parameter a
bound
, are extracted by least-
squares fitting of a biexponential decay at each pixel
11,24–26
.
Drawbacks of this method of analysis have been identified, such
as the computational burden of fitting 10
4
–10
5
independent decay
curves in a single image, difficulties in resolving decay
components with closely spaced lifetimes and correlations
between the fit parameters
52
. Other groups have therefore
focussed on developing novel analytical techniques. For
example, Yaseen et al.
53
recently applied global analysis to the
application of NAD(P)H FLIM of the rat cortex. In this approach,
the lifetimes of four decay components were shared across
the image and a novel computational algorithm recovered the
optimum amplitudes of each species to describe the fluorescence
decay at each pixel. Interestingly, two enzyme-bound components
were identified in the tissue with lifetimes of 1.7 and 3.2 ns,
perhaps corresponding to enzyme-bound NADH and NADPH.
Another approach to which our conclusions may be strongly
applicable is the phasor method developed by Digman et al.
52
Here, the real and imaginary parts of the Fourier transform of the
fluorescence decay at each pixel of a FLIM image define the
coordinates of a point in a two-dimensional phase space.
The relative abundance of two or more fluorescent species with
different lifetimes can then be inferred from the location of the
pixels in the phasor plot. Application of this ‘fit-free’ approach to
separating NADH and NADPH fluorescence will be the subject of
further work.
Altogether, we have shown that the fluorescence lifetime
characteristics of NAD(P)H in live cells and tissues can be used to
discriminate between NADH and NADPH fluorescence, provid-
ing, for the first time, a biochemical framework for interpretation
of NAD(P)H FLIM studies. Such a technique permits the
separation of NADH and NADPH redox signalling without
disrupting the sample on the addition of external probes
54,55
,
allowing complex tissue preparations to be investigated. The
approach revealed previously unknown cellular metabolic
specializations in the mammalian cochlea, highlighting a
subpopulation of cells characterized by high NADPH levels,
opening up new avenues of research to understand the functional
significance of redox pathways with respect to the physiological
roles of these cells.
Methods
Cell culture. HEK293 cells were obtained from the American Type Culture
Collection and grown in advanced Dulbecco’s modified Eagle Medium (Gibco)
supplemented with 10% fetal bovine serum, 2 mM GlutaMAX, 100 U ml
1
penicillin and 100 mgml
1
streptomycin (Gibco). Production of the NADK þ and
NADK cell lines has been reported previously
29
. These cultures were
additionally supplemented with 0.1 mg ml
1
G-418 selective antibiotic (Gibco).
All cells were grown as monolayers in sterile 75 cm
2
tissue culture flasks (Thermo
Scientific Nunc) in a 37 °C, 5% CO
2
incubator.
Cochlea explant cultures
. Cochlear coils were isolated from male and female
post-natal day 2–3 Sprague Dawley rats as previously described
56
. Briefly, auditory
bullae were removed and transferred into Medium 199 with Hank’s balanced salts
(Life Technologies). The cartilaginous wall of the bulla was opened and the whole
cochlea extracted. The stria vascularis and Reissner’s membrane were removed and
the cochlea cut into three coils. The cochlear coils were placed onto Cell-Tak cell
and tissue adhesive (BD Biosciences)-coated dishes (MatTek). For coating, cell
adhesive was diluted to 70 mgml
1
in 0.1 mM NaHCO
3
. The cochlear explants
were incubated overnight in DMEM/F12 (Gibco), supplemented with 1% fetal
bovine serum (Life Technologies) in a 37 °C, 5% CO
2
incubator. The isolation was
performed in accordance with the United Kingdom Animals (Scientific
Procedures) Act of 1986 and in compliance with the Biological Services
Management Group and the Biological Services Ethical Committee, University
College London.
Live-cell microscopy
. On the microscope stage, coverslips containing 300,000 cells
were maintained at 37 °C in a metal ring and bathed in DMEM solution at pH 7.4
containing 10 mM HEPES and 2 mM GlutaMAX. Glucose (25 mM) was present
under contro l conditions and during EGCG treatment (100 mM in cell lines,
200 mM for cochlea). Pharmacological perturbations to metabolism were applied by
the dropwise addition of working concentrations of each compound, diluted from
stock solutions in DMEM recording medium.
NAD(P)H FLIM was performed on an upright LSM 510 microscope (Carl
Zeiss) with a 1.0 NA 40 water-dipping objective using a 650-nm short-pass
dichroic and 460
±
25 nm emission filter. Two-photon excitation was provided by a
Chameleon (Coherent) Ti:sapphire laser tuned to 700 nm, with on-sample powers
kept below 10 mW. Spectral controls (see Supplementary Fig. 4 and Supplementary
Note 5) confirmed the NAD(P)H specificity of this excitation wavelength and
emission filtering. Photodamage controls (see Supplementary Tables 6 and 7 and
Supplementary Note 6) demonstrated that FLIM parameters were not varying over
the course of imaging. Emission events were registered by an external detector
(HPM-100, Becker & Hickl) attached to a commercial time-correlated single
photon counting electronics module (SPC-830, Becker & Hickl) contained on a
PCI board in a desktop computer. Scanning was performed continuously for 4 min
with a pixel dwell time of 1.6 ms. To identify mitochondrial and cytosolic regions,
the mitochondrially targeted fluorescent dye tetramethylrhodamine methyl ester
(TMRM) was added to the recording medium, at a final concentra tion of 25 nM,
20 min before imaging. TMRM fluorescence was collected for a 10-s burst using a
610
±
30 nm emission filter. Excitation was provided at the same wavelength as
NAD(P)H to avoid possible chromatic aberration. The 585
±
15 nm emission
spectrum of TMRM ensured its fluorescence did not contaminate the NAD(P)H
FLIM images.
NAD(P)H fluorescence intensity time series and MCB imaging were performed
on an inverted LSM 510 laser scanning confocal microscope (Carl Zeiss) with
351 nm illumination from an argon ion laser (Coherent Enterprise UV). MCB
(30 min loading at 50 mM) and NAD(P)H fluorescence were observed using a
351-nm long-pass dichroic and 460
±
25 nm band-pass emission filter with a 40,
1.3 NA quartz oil immersion objective. Images (12-bit 512 512) were obtained
with a pixel dwell time of 1.6 ms. Time series measurements were obtained at 1 min
intervals. To reduce noise, the image recorded at each time point was a mean of
four consecutive scans. Fluorescence intensity levels were extracted using ImageJ
(NIH).
Metabolic controls
. For the measurement of oxygen consumption, cells were
trypsinized and resuspended at B1 million cells per ml in DMEM buffered with
10 mM HEPES and supplemented with 25 mM glucose and 2 mM GlutaMAX.
Respiration rates were measured in triplicate at 37 °C with the high-resolution
Oxygraph (Oroboros Oxygraph-2k). State 4 respiration values were obtained in the
presence of 2.5 mM oligomycin, maximal oxidative capacities were determined in
the presence of 2 mM FCCP and non-mitochondrial background oxygen con-
sumption was determined in the presence of 2.5 mM antimycin A. Lactate release
rates were measured in triplicate by removing a sample of serum-free DMEM from
70% confluent cell cultures at 1 h intervals for 5 h total and determining the
concentration of lactate present using a commercially available plate-reader assay
(Sigma Aldrich) in absorption mode. For normalization of both oxygen
consumption and lactate release rates, final cell counts were performed using a
haemocytometer.
FLIM data analysis
. Following 5 5 binning of photon counts at each pixel,
fluorescence decay curves of the form
ItðÞ¼Z þ I
0
1 a
bound
ðÞexp
t
t
free
þ a
bound
exp
t
t
bound
ð2Þ
were fit to the FLIM images using iterative reconvolution in SPCImage (Becker &
Hickl), where Z allows for time-uncorrelated background noise. The instrument
response function of the FLIM system was obtained by measuring the fluorescence
decay profile of second harmonic generation by a potassium dihydrogen phosphate
(KDP) crystal at 920 nm, grown by leaving a molar solution of KDP in water on a
coverslip to evaporate. Matrices of the fit parameters t
free
, a
bound
and t
bound
, along
NATURE COMMUNICATIONS | DOI: 10.1038/ncomms4936 ARTICLE
NATURE COMMUNICATIONS | 5:3936 | DOI: 10.1038/ncomms4936 | www.nature.com/naturecommunications 7
& 2014 Macmillan Publishers Limited. All rights reserved.
with the total photons counted at each pixel, were exported from SPCImage.
In MATLAB (The Mathworks), a 16-bit grayscale image was produced for each
parameter matrix in which the intensity of each pixel was proportional to the
parameter value at that location. ImageJ was then used to measure the grayscale
intensity in the parameter images and the values obtained converted back to
parameter values using the scaling factors applied in their production. Masks
identifying the location of mitochondrial and cytosolic pixels were created by
importing images of the TMRM distribution in the cell and a nuclear mask was
defined by hand, allowing the parameters describing the fluorescence decay of
NAD(P)H in each of these regions to be extracted separately (Supplementary
Fig. 2a).
Statistical analysis
. NAD(P)H fluorescence lifetime parameters obtained using
FLIM are reported as a mean over at least three regions of at least three separate
cultures. Uncertainties in these values were taken as the s.d. of the measurements.
Differences between the lifetime parameters measured under different conditions
were tested for statistical significance (Po0.05) using a two-tailed Student’s t-test.
Statistically significant differences between data sets normalized to be expressed in
relative arbitrary units (fluorescence intensity, NADH/NADPH concentration)
were assessed using a Wilcoxon signed-rank test. For the estimation of uncer-
tainties in the predictions of the numerical model, the standard formula for the
calculation of the error s
2
z
in an arbitrary function Z ¼ Z(A, B, C,...) was applied,
s
2
z
¼
@Z
@A
2
s
2
A
þ
@Z
@B
2
s
2
B
þ
@Z
@C
2
s
2
C
::: ð3Þ
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Acknowledgements
We thank Dr Will Kotiadis for assistance with respirometry measurements, Dr Zhi Yao
for assistance with the lactate release assay and Dr Laura Osellame for proofreading
of the manuscript. Action on Hearing Loss supported the work of Z.F.M. and J.E.G.
G.S. was funded by Parkinson’s UK, British Heart Foundation, Wellcome Trust-UCL
Therapeutic Innovation Fund, Telethon (Italy, GEP1206) and the Italian Association for
Cancer Research (AIRC). M.R.D. acknowledges strategic fu nding from the Wellcome
Trust/MRC Joint Call in Neurodegeneration Award (WT089698). T.S.B. is grateful
for the award of an EPSRC studentship and the inaugural Professor Anne Warner
postdoctoral fellowship through the CoMPLEX Doctoral Training Centre at UCL.
Authors contributions
M.Z. provided the NADK þ and NADK– cell lines. J.E.G. prepared cochlea cultures and
oversaw their imaging. Z.F.M., G.S. and M.R.D. performed preliminary experiments.
T.S.B. performed the experiments, data analysis and modelling. A.J.B., G.S. and M.R.D.
supervised the work. T.S.B., G.S. and M.R.D. wrote the manuscript.
Additional information
Supplementary Information accompanies this paper at http://www.nature.com/
naturecommunications
Competing financial interests: The authors declare no competing financial interests.
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reprintsandpermissions/
How to cite this article: Blacker, T. S. et al. Separating NADH and NADPH fluorescence
in live cells and tissues using FLIM. Nat. Commun. 5:3936 doi: 10.1038/ncomms4936
(2014).
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