NAD(P)H fluorescence lifetime imaging of glucose stimulated MIN6
cells
Raluca Niesner1,2, Bülent Peker1, Stefan Quentmeier1, Stefan Denicke1, Ingo Rustenbeck3 and KarlHeinz Gericke1*
1
Institute of Physical and Theoretical Chemistry, Technical University of Braunschweig, HansSommer Straße 10, D-38106 Braunschweig, Germany
2
Helmholtz Centre for Infection Research, Inhoffenstraße 7, D-38124 Braunschweig, Germany
3
Institute of Pharmacology and Toxicology, Technical University of Braunschweig, Mendelssohn
Straße 2, D-38106 Braunschweig, Germany
Short title:
NAD(P)H time-resolved imaging of stimulated MIN6 cells
Correspondence address:
Karl-Heinz Gericke, Prof. Dr.
Technical University Braunschweig
Institute for Physical and Theoretical Chemistry
Hans-Sommer Str. 10
D-38106 Braunschweig
Germany
phone: +49 (531) 391 5325/26
fax: +49 (531) 391 5396
e-mail: [email protected]
Abstract
The coenzymes nicotine amide adenosine dinucleotide (NADH) and nicotine amide adenosine
dinucleotide phosphate (NADPH), hereafter NAD(P)H, have a particular relevance for the
biosciences since they are ubiquitous electron carrier in vital cellular processes. Especially the fact,
that only their reduced forms fluoresce, makes these coenzymes the ideal signal mediators for
monitoring the redox metabolic activity within cells, i.e. the redox equilibrium between reduced
NAD(P)H and oxidised NAD(P). Up to now, most techniques based on endogenous NAD(P)H
fluorescence directly calculate the relative concentration of NAD(P)H from the total NAD(P)H
fluorescence signal, i.e. steady-state techniques. However, they ignore the fact that cellular
NAD(P)H is either free (metabolically inactive) or bound to enzymes, if it participates to vital redox
processes. Since these two NAD(P)H states have different fluorescence properties (lifetimes and,
thus, quantum yields), the total NAD(P)H fluorescence signal is strongly influenced by their
relative concentrations. Neglecting this dependence leads to erroneous interpretations of the level of
the total NAD(P)H fluorescence signal and, thus, to erroneous relative NAD(P)H and NAD(P)
concentrations. We demonstrate for the first time that only biexponential NAD(P)H fluorescence
lifetime imaging (FLIM) allows an accurate quantitative evaluation of the contributions of the
equilibria NAD(P)(ox)  NAD(P)H(red) and NAD(P)H(free)  NAD(P)H(enzyme-bound) to the
total NAD(P)H fluorescence signal and, thus, a precise evaluation of the cellular metabolic activity.
Moreover, due to the subcellular resolution of our method based on two-photon microscopy, this
quantification resembles the cellular heterogeneity, i.e. allows the quantification of the relative
coenzyme concentrations in different regions of the cell (nuclei and organelles within the
cytoplasm). Although demonstrative experiments have been performed on MIN6 cells (mutated
insulin-producing -cells) stimulated with glucose, our method is generally applicable to monitor
the cellular metabolic activity of all types of biological systems.
Keywords: endogenous NAD(P)H fluorescence, biexponential NAD(P)H fluorescence lifetime
imaging, glucose stimulation, MIN6 cells
1
Introduction
The coenzymes nicotine amide adenosine dinucleotide NADH and nicotine amide adenosine
dinucleotide phosphate NADPH (hereafter NAD(P)H) are ubiquitous electron-carrier in central
metabolic redox processes, e.g. ATP production and reductive biosyntheses of macromolecules
within the cell [1]. Considering the ability of their reduced forms to fluoresce, it is straightforward
that these coenzymes are particularly appropriate as signal mediators for monitoring vital cellular
processes under non-invasive genuine conditions. This fact is mirrored in the multitude of research
works, which employ (bulk and spatially resolved) NAD(P)H-fluorescence techniques to 3D-image
tissue morphology as well as to study cellular function and cellular function disorders [1-10].
Spatially resolved (imaging) techniques are naturally of greater relevance for the biosciences since
they resemble the cellular heterogeneity and, thus, provide more detailed information [5-10].
Furthermore, it is well known that the laser scanning fluorescence microscopy based on two-photon
excitation is more appropriate for investigations in thick highly-scattering biological media, e.g.
tissue or even organs, than the standard confocal microscopy based on one-photon excitation [7,1113]. The reasons therefore are the intrinsic 3D resolution, which allows optical sectioning of the
sample and leads to a low photodamage and photobleaching outside the focal plane, and a large
penetration depth in tissue due to the use of near infrared (NIR) illumination [14].
Particularly the quantification of the redox equilibrium between the reduced (fluorescent) NAD(P)H
and the oxidised (non-fluorescent) NAD(P), i.e. the cellular redox metabolic activity, is a central
aim for all the techniques based on endogenous NAD(P)H fluorescence since this is generally
relevant for the main bioscientific branches. Up to now, most techniques used for this purpose are
steady-state techniques, in which the relative NAD(P)H concentration is directly calculated from
the total NAD(P)H fluorescence signal [5,6,8,9,13]. However, these methods ignore the fact, that
there are two states of the reduced (fluorescing) coenzymes within the cell – free (metabolically
inactive) and bound to enzymes during redox processes. These two NAD(P)H states are
characterised by specific fluorescence properties, i.e. overlapping emission spectra but strongly
differing fluorescence lifetimes and, thus, very different fluorescence quantum yields [12,15-18].
Consequently, the relative concentrations of the free and enzyme-bound NAD(P)H, respectively,
dramatically influence the cellular NAD(P)H fluorescence quantum yield averaged over all
NAD(P)H states and, thus, the level of the total NAD(P)H fluorescence signal. Since steady-state
techniques cannot even distinguish between these two states, the NAD(P)H concentration calculated
thereby is imprecise. The few works, which take into account, the existence of free and enzymebound NAD(P)H, usually correct the relative NAD(P)H concentration attained in steady-state
experiments by dividing it by the lifetime ratio of the free and enzyme-bound NAD(P)H [19]. This
correction is inaccurate because it does not take into account either the heterogeneously distributed
2
concentrations of the cellular free and bound NAD(P)H or the fact that the fluorescence lifetime of
the enzyme-bound NAD(P)H strongly depends on the enzyme, to which NAD(P)H is bound to
[12,18].
In this work, we accurately quantify the contributions of the equilibria NAD(P) (ox)  NAD(P)H
(red) and NAD(P)H (free)  NAD(P)H (enzyme-bound), respectively, to the total NAD(P)H
fluorescence signal within cells by means of biexponential NAD(P)H fluorescence lifetime
imaging. Thus, we are for the first time able to precisely determine the cellular NAD(P)H
concentration and, thus, the redox metabolic activity within cells. The employed FLIM technique is
particularly adequate for such measurements since it allows to quantitatively resolve between free
and enzyme-bound NAD(P)H and, thus, to exactly determine the averaged NAD(P)H fluorescence
quantum yield. Moreover, due to the subcellular resolution of the employed technique based on
two-photon microscopy, this quantification resembles the cellular heterogeneity, i.e. it allows the
quantification of the relative NAD(P) and NAD(P)H concentrations in different regions of the cell
like nuclei and organelles within the cytoplasm.
The biological model, on which we performed demonstrative experiments, was the suspension of
MIN6 cells (mutated insulin producing -cells) stimulated with 20 mmol/L solution of glucose.
However, the results of our experiments show that biexponential NAD(P)H FLIM is universally
applicable to monitor the cellular metabolic activity with high accuracy and at a high spatial
resolution.
Theoretical approaches and data analysis
We consider that the fluorescence emitted by MIN6 cells under the given experimental conditions
(two-photon excitation at 760 nm) originates only from the reduced forms of the coenzymes
nicotine-amide adenine dinucleotide and nicotine-amide adenine dinucleotide phosphate (NADH
and NADPH) as we will show later. The oxidised forms of these coenzymes (NAD and NADP) are
non-fluorescent. The concentrations of the oxidised and reduced forms within the cell are related by
redox reactions catalysed by substrate- and coenzyme-specific enzymes:
NAD(P) + substrate (red)  substrate (ox) + NAD(P)H.
Steady-state approach: The total cellular NAD(P)H fluorescence signal F after two-photon
excitation is given by:
F     c  2    c
(1)
with  the two-photon excitation cross-section,  the fluorescence quantum yield, c the
concentration of the chromophores and  the excitation photon flux at the sample.
3
When MIN6 cells are stimulated for a few minutes with glucose media (here, 20 mmol/L) after they
have been kept on media containing 0 mmol/L glucose, the NAD(P)H fluorescence signal F
increases [20,21]. The ratio between the fluorescence signal of the cells stimulated with glucose
(F20) and that of the cells kept in 0 mmol/L glucose buffer (F0) is given by:
F 20  20 c 20
 0  0 .
F0
 c
(2)
We assume that the proportionality constant in Eq. 1 does not change because comparative
measurements were performed at the same laser power, i.e. at the same photon flux , the
stimulation with glucose can not influence the symmetry of the electronic states in the coenzyme
molecules, which would lead to a modification of the two-photon excitation cross-section  and the
signal of all possible states of NAD(P)H is detected in the same way no matter of the stimulation of
the cells.
Concluding, the increase of the total NAD(P)H fluorescence signal under stimulation with glucose
is to be explained either by an increase of the NAD(P)H concentration as the consequence of a shift
in the equilibrium NAD(P)  NAD(P)H or by an increase of the (averaged) NAD(P)H fluorescence
quantum yield  or by both of them.
All redox reactions based on the considered coenzymes are steps of vital cellular processes and are
catalysed by enzymes, which bind to the coenzymes. Thus, both free NAD(P)H, which does not
participate to any vital process, and enzyme-bound NAD(P)H, which takes part to redox reactions,
are present and fluoresce within the cell. Consequently, the total NAD(P)H fluorescence signal is
given by:
F  F1  F2  1  c1  2  c2 .
(3)
Index 1 indicates inhere the free NAD(P)H and index 2 the enzyme-bound NAD(P)H. We assume
that the two-photon excitation cross-sections of the NAD(P)H chromophores is the same for the free
and enzyme-bound state, respectively, and that both states are identically excited and detected.
Thus, the ratio F20/F0 is given by:



F 20 c120  c220  220 120
 0
.
F0
c1  c20  20 10

(4)
Results of FLIM experiments on MIN6 cells indicate that the fluorescence lifetimes and  of the
free and enzyme-bound NAD(P)H, respectively, do not change under stimulation with glucose (data
shown in the following sections). Since the fluorescence quantum yield is defined by     k f ,
where the fluorescence rate kf is independent of the molecular environment, we conclude that
120  10  1 , 220  20  2 and 2 1   2  1 . Thus, Eq. (4) becomes:
4
F 20 c120  c220   2  1 
.
 0
F0
c1  c20   2  1 
(5)
Since free and enzyme-bound NAD(P)H, respectively, have different fluorescence properties as
shown above, a modification of their relative concentrations lead to a modification of the averaged
fluorescence quantum yield  of the cellular NAD(P)H.
Time-dependence approach: The presence of both free and enzyme-bound NAD(P)H within the
cell justifies the assumption that the time-dependent NAD(P)H fluorescence signal F(t) is best
described by a biexponential function:
F (t )  a1  e
 t
1
 a2  e
t
2
,
(6)
where the first exponential function describes the fluorescence decay of the free NAD(P)H and the
second function that of the enzyme-bound NAD(P)H. By integrating these fluorescence decays over
time, we obtain the total fluorescence signal F1  a1  1 for the free NAD(P)H and F2  a2  2 for
the enzyme-bound NAD(P)H. The parameters a1, a2, 1 and 2 are determined from biexponential
FLIM data based on the NAD(P)H fluorescence of MIN6 cells.
The relations between the parameters ai and the concentrations ci (i = 1, 2) are:
Fi  ai   i  i  ci
 ai 
i  ci
 k f  ci .
i
(7)
Thus, the ratio Y = a2/(a1 + a2) calculated from the FLIM data is equal to the relative concentration
of the enzyme-bound NAD(P)H: c2/(c1 + c2). The sum c1 + c2 represents the total NAD(P)H
concentration c within the cell.
Combining the result of the steady-state approach with that of the time-dependence approach, the
ratio F20/F0 (Eq. 5) can be further expressed as:
F 20 c120  c220   2  1  c 20  c220   2  1  1 c 20 1  Y 20   2  1  1
.
 0
 0
 0 
F0
c1  c20   2  1 
c  c20   2  1  1
c 1  Y 0   2  1  1
(8)
By comparing Eq. (8) with Eq. (2), we obtain:
 20 1  Y 20   2  1  1

.
 0 1  Y 0   2  1  1
(9)
Consequently, the results of biexponential FLIM experiments based on the NAD(P)H fluorescence
directly indicate in which proportion the shift in the equilibrium NAD(P)  NAD(P)H and the
modification of the relative concentrations of free and enzyme-bound NAD(P)H contribute to the
increase of the total NAD(P)H fluorescence signal F in (MIN6) cells under stimulation with
glucose. Note that a shift in the equilibrium NAD(P)  NAD(P)H leads to a change in the total
5
NAD(P)H concentration c, whereas a modification of the relative concentrations of the free and
enzyme-bound NAD(P)H leads to a change in the averaged fluorescence quantum yield .
Total endogenous NAD(P)H fluorescence of MIN6 cells
It has been already demonstrated that the endogenous fluorescence of cells, which are excited at
approx. 750 nm (two-photon excitation), mainly originates from the coenzymes NADH and
NADPH [22]. The fluorescence of these coenzymes can be easily separated from that of the only
possible interfering chromophores, i.e. flavine adenosine dinucleotide FAD, by using appropriate
detection filters.
Hence, the total fluorescence signal of MIN6 cells excited at 760 nm and detected through an
NAD(P)H interference filter (460±20 nm) stems from (free and enzyme-bound) NAD(P)H. The
fact, that the signal is low in the cell nuclei but very high in small organelles in the cytoplasm, i.e.
mitochondria, confirms this assumption (Fig. 1).
After stimulating the MIN6 cells with 20 mmol/L glucose media an increase of the total NAD(P)H
fluorescence signal F of 33.2±2.5 % is observed (Fig. 1). The fluorescence signal returns to the
previous level after removing the stimulus. This corresponds to a signal decrease of 32.4±2.1 %. In
all cases the percentage results refer to
F 20  F 0
 100 .
F0
Insert Fig. 1
It is noteworthy that the fluorescence increase under stimulation with 20 mmol/L glucose within the
cell nuclei is significantly lower (12.6±2.1 %) than in cytoplasm (47.8±10.1 %). Furthermore, the
fluorescence increase in different regions in cytosol strongly varies: increases between 29.4 % and
61.5 % have been measured.
Concluding, steady-state measurements allow us to quantify the increase of the total NAD(P)H
fluorescence signal F in MIN6 cells under stimulation with glucose, which is obviously related to
an increased metabolic activity in mitochondria (in cytoplasm). However, they do not give any clue
about the biomolecular reasons for this signal increase: a shift in the redox equilibrium NAD(P) 
NAD(P)H or an increased NAD(P)H fluorescence quantum yield .
Biexponential NAD(P)H-FLIM
As already mentioned in the introduction, biexponential NAD(P)H fluorescence lifetime imaging
(NAD(P)H-FLIM) allows the differentiation between free (inactive) and enzyme-bound (active)
NAD(P)H in each pixel of the time-resolved fluorescence image. A further result of NAD(P)HFLIM is the map of the relative contribution of the enzyme-bound NAD(P)H to the total NAD(P)H
signal Y·100 = a2·100 /(a1 + a2), i.e. ratio map.
6
Insert Fig. 2
FLIM experiments on MIN6 cells confirm the differentiation between free and bound NAD(P)H
based on the fluorescence lifetime (Fig. 2a and b). The average value over 18 lifetime images (42
cells) of free NAD(P)H is 450±40 ps, which is in good agreement with the lifetime values of
NADH and NADPH measured in buffer solution, i.e. 440 ps and 412 ps, respectively [12]. As far as
the lifetime of the enzyme-bound NAD(P)H is concerned, the average value is 2980±140 ps. As
expected, the fluorescence lifetime 2 of bound NAD(P)H is significantly longer than that of the
free NAD(P)H. Furthermore, the strong dependence of 2 on the enzyme, to which NAD(P)H is
bound to, resembles in the wide distribution (approx. 1300 ps) of this lifetime within the cell (see
2-histogram in Fig. 2d). The ratio map in Fig. 2c indicate a high metabolic activity of the MIN6
cells (98±1 % contribution of the enzyme-bound NAD(P)H).
The differentiation between free and enzyme-bound NAD(P)H is the necessary precondition to
answer the question about the biomolecular reasons of the fluorescence signal increase in MIN6
cells under stimulation with glucose.
NAD(P)H-FLIM study of glucose stimulation
Also in time-resolved experiments, i.e. biexponential NAD(P)H-FLIM experiments, the total
fluorescence signal F of MIN6 cells show the same behaviour under stimulation with 20 mmol/L
glucose as in steady-state experiments. An increase of 32.8±2.5 % was observed under stimulation,
whereas the decrease after removing the stimulus was 32.1±2.4 %.
As far as the fluorescence lifetime of both free and enzyme-bound NAD(P)H is concerned, we
could not observe any modification under stimulation with glucose. In glucose free media we
measured 450±40 ps for free NAD(P)H and 2980±140 ps for bound NAD(P)H, while in 20 mmol/L
glucose media 440±60 ps for free NAD(P)H and 3020±120 ps for bound NAD(P)H were
determined. Considering these results, we can assert that the fluorescence quantum yields 1 and 2
of the free and bound NAD(P)H, respectively, are independent of the glucose stimulation. Thus, the
NAD(P)H fluorescence quantum yield  varies only when the relative concentrations of the free and
enzyme-bound NAD(P)H change.
Modifications of the relative (free and enzyme-bound) NAD(P)H concentrations are visible in the
ratio map Y·100 = a2·100/(a1 + a2), which corresponds to c2·100/(c1 + c2). As shown in the
theoretical section, in order to facilitate the comparison between steady-state and time-resolved
data, we calculate the ratio 0/20 of the NAD(P)H fluorescence quantum yield from the relative
concentrations ratio Y0/Y20. 0/20 naturally resembles only modifications of the total fluorescence
7
signal F caused by a modification of the NAD(P)H quantum yield  and not by a shift in the redox
equilibrium NAD(P)  NAD(P)H.
Insert Fig. 3
Under stimulation with 20 mmol/L glucose, we observed an increase in the proportional image
calculated after: 1  Y   2 1  1 (Fig. 3a). This increase is obviously related to an increase of the
relative concentration of the enzyme-bound NAD(P)H. A corresponding decrease of this
concentration was observed after removing the stimulus (Fig. 3b).
Statistics over 40 cells revealed that the average cellular increase of the total fluorescence signal F
due to modifications of  amounts to 23.6±2.1 % under stimulation with 20 mmol/L glucose, while
the decrease after removing the stimulus is 23.3±1.5 %.
Considering Eq. (2), the average increase of the total fluorescence signal (32.4±2.5 %) under
stimulation with glucose is caused both by a modification of the NAD(P)H fluorescence quantum
yield  (23.5±1.8 %) and by a shift of the redox equilibrium NAD(P)  NAD(P)H (7.3±3.2 %).
Within the nuclei, the increase of the fluorescence signal (12.6±2.1 %) is due only to an increase of
quantum yield (13.2±5.3 %). No measurable shift in the redox equilibrium NAD(P)  NAD(P)H
was observed. In contrast, the increase of fluorescence signal in cytoplasm (47.8±10.1 %) is due to
both the redox shift (12.9±6.3 %) and the modification of the fluorescence quantum yield 
(30.9±6.6 %).
Summary
Most of the steady-state experiments, in which the cellular NAD(P)H concentration is determined,
start from the assumption that an increase of the total NAD(P)H fluorescence signal is necessarily
caused by a shift in the redox equilibrium NAD(P)  NAD(P)H, since it is well known that
NAD(P) is non-fluorescent, while NAD(P)H fluoresces. However, this assumption neglects the fact
that fluorescent NAD(P)H within cells appears both as free (metabolically inactive) and as enzymebound (metabolically active). These NAD(P)H states are very different as far as their fluorescence
properties are concerned, i.e. they have different fluorescence lifetimes and, thus, different
fluorescence quantum yields. Consequently, the cellular NAD(P)H concentration calculated from
the increase of the total NAD(P)H fluorescence signal is imprecise.
We demonstrate that the cellular NAD(P)H concentration can quantitatively be determined only by
means of biexponential NAD(P)H-FLIM (time-resolved technique), since this method provides
information about the biomolecular reasons of the increase in the total NAD(P)H fluorescence
signal. This means, it accurately quantifies the contributions of the shift in the redox equilibrium
NAD(P)  NAD(P)H and of the ratio free NAD(P)H to enzyme-bound NAD(P)H in each pixel of
8
the image. The main results of the steady-state as well as of the biexponential NAD(P)H-FLIM
experiments on MIN6 cells stimulated with glucose are summarised in Table 1.
Insert Table 1
Due to the subcellular spatial resolution of our method, we are able to accurately determine the
increase of the total NAD(P)H fluorescence signal, of the NAD(P)H concentration and of its
fluorescence quantum yield under stimulation with glucose in nuclei regions and in cytoplasmic
regions with a high content of mitochondria.
Concluding, by means of biexponential NAD(P)H FLIM we were for the first time able to
quantitatively determine the NAD(P)H redox metabolic activity of cells with a high spatial (and
time) resolution.
Methods
All experiments were carried out using a specialised two-photon laser-scanning microscope based
on a tunable (720 to 920 nm) Ti:Sa laser (MaiTai, Spectra Physics, Darmstadt, Germany) and on a
commercial scan-head (TriMScope, LaVision BioTec, Bielefeld, Germany), which allows multibeam (up to 64 beam lets) scanning of the sample without cross-talk between neighboring beam
lets. The multi-beam scanning permits a speed-up of data acquisition proportional to the splitting of
the main laser beam. The excitation beam is focused into the sample by a 20 objective lens with
NA = 0.95 and a working distance of 2 mm (Olympus, Hamburg, Germany). In steady-state
experiments we employed as detection unit a CCD camera (Sensicam QE, LaVision, Göttingen,
Germany). In FLIM experiments, i.e. time-resolved experiments, the fluorescence signal was
detected by using an intensified CCD camera at 500 ps time-gate (PicoStar, LaVision). The
NAD(P)H fluorescence was detected through an interference filter (HQ 460±20 nm, AHF,
Germany).
The spatial resolution in the steady-state experiments was 350 nm (lateral) and 1.4 µm (axial). In
the time-resolved experiments, we measured a worse lateral spatial resolution of 550 nm. The
spatial resolution was determined by using 100 nm fluorescing polystyrene beads.
For all experiments, mutated insulin producing MIN6 cells were seeded on sterile cover slips, which
were used to air-tight close a self-designed flow chamber for microscopy. Carbogen (95% CO2 and
5% O2) saturated Krebs-Ringer-Hepes buffer with 0mmol/L and 20 mmol/L glucose, respectively,
was used as media for the experiments. All experiments were performed at 37°C.
Acknowledgement
We acknowledge the Bundesministerium für Bildung und Forschung for financial support under
grant 0313412C (Bioprofile Hannover/Braunschweig/Göttingen).
9
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10
FIGURE CAPTIONS
Figure 1: a) and b) Increase and decrease of the NAD(P)H fluorescence signal F in MIN6 cells
during stimulation with 20 mmol/L glucose and after removing the stimulus, respectively. The time
lapse between two consecutive (shown) images was 10 minutes. Excitation wavelength  = 760 nm,
emission filter 460±20 nm. Note that the signal increase within the nuclei is much lower than in the
cytoplasm (mitochondria).
Figure 2: Result of biexponential NAD(P)H fluorescence lifetime imaging on MIN6 cells: a)
fluorescence lifetime map (1-map) of the free NAD(P)H; b) fluorescence lifetime map (2-map) of
the enzyme-bound NAD(P)H, which participates to redox processes catalysed by enzymes to which
it is bound to; c) contribution of the bound NAD(P)H to the total fluorescence signal as well as
relative concentration of the bound NAD(P)H to the total NAD(P)H concentration, i.e. ratio Y·100
= a2·100/(a1 + a2) map equivalent to the enzyme-bound NAD(P)H relative concentration c2·100/(c1
+ c2) map. d) Distributions of 1 and 2 in the images a) and b). The values of 1 and 2 confirm that
free and bound NAD(P)H are temporally resolved. Time resolution is approx. 10 ps.
Figure 3: a) Increase and b) decrease of 1  Y   2 1  1 in the corresponding images under
glucose stimulation and after removing the stimulus, respectively. The expression 1  Y   2 1  1
is proportional to the NAD(P)H fluorescence quantum yield  under the given conditions and is not
influenced by modifications in the redox equilibrium NAD(P)  NAD(P)H.
TABLE LEGEND
Table 1: Increase of the total NAD(P)H fluorescence signal F averaged over whole MIN6 cells,
nuclei regions as wells as cytoplasmic regions under glucose stimulation. Contributions of the
NAD(P)H fluorescence quantum yield  as well as of the redox equilibrium NAD(P)  NAD(P)H
to these modifications of the fluorescence signal F.
11
(F20-F0)/F0
(F20-F0)/F0
(steady-state)
(time-resoved)
cell
32.8±2.2 %
nucleus
cytoplasm
(20-0)/0
(c20-c0)/c0
32.4±2.5 %
23.5±1.9 %
7.3±2.1 %
12.6±2.1 %
–
13.2±5.3 %
47.8±10.1 %
–
30.9±6.6 %
0.5±5.0 %
(approx. 0 %)
12.9±6.3 %
12
a
b
10
µm
Figure 1
0
mM
20
mM
0
mM
13
a
b
10 µm
[p
s]
[p
s]
d
c
%
free NAD(P)H
(463 ps, FWHM = 288 ps)
enzyme-bound NAD(P)H
(2864 ps, FWHM = 1341 ps)
1,0
0,9
0,8
rel. occurance
0,7
0,6
0,5
0,4
0,3
0,2
0,1
0,0
0
500
1000 1500 2000 2500 3000 3500 4000 4500 5000 5500 6000
 [ps]
Figure 2
14
0 mM
a
20 mM
a.u.
10 µm
20 mM
b
0 mM
a.u.
Figure 3
15