IEEE JOURNAL OF SELECTED TOPICS IN QUANTUM ELECTRONICS, VOL. 13, NO. 6, NOVEMBER/DECEMBER 2007
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Fluorescence Anisotropy of Cellular NADH as a Tool
to Study Different Metabolic Properties of Human
Melanocytes and Melanoma Cells
Ye Yuan, Yanfang Li, Brent D. Cameron, and Patricia Relue
Abstract—In this study, we wanted to see if fluorescence
anisotropy could be used to detect changes in metabolism in cells
with significant light scattering and absorption properties. Fluorescence anisotropy measurements of nicotinamide adenine dinucleotide (NADH) were performed with human melanocytes and
melanoma cell lines. To demonstrate the feasibility of using fluorescence anisotropy for detecting metabolic changes, the electron
transport chain was blocked using rotenone, inducing an accumulation of intracellular NADH. Total fluorescence increased in all
cells as a result of rotenone treatment. Fluorescence anisotropy
decreased in the rotenone-treated cells relative to the controls,
suggesting an increased ratio of free to protein-bound NADH in
the treated cells. In general, the fluorescence anisotropy of the
melanocytes was significantly higher than that of the melanoma
cell lines. Reflectance spectroscopy showed that the differences in
fluorescence anisotropy between the cell types were not due to
differences in scattering and absorption properties. Intrinsic cellular NADH fluorescence was experimentally extracted by ratioing
polarized fluorescence to polarized reflectance. NADH binding,
measured as the ratio of fluorescence intensity at 430 and 465
nm, showed more protein-bound NADH in the melanocytes than
in the melanoma cells, consistent with the fluorescence anisotropy
measurements.
Index Terms—Cancer, fluorescence spectroscopy, metabolism,
polarized fluorescence, reflectance spectroscopy.
Fig. 1. Summary of NADH binding with associated enzymes involved in aerobic and anaerobic metabolism. Glucose conversion to pyruvate is common
to both metabolic pathways. From pyruvate, anaerobic metabolism yields lactate; two NADH binding sites exist within anaerobic metabolism. For aerobic
metabolism, pyruvate enters the mitochondria where energy is produced via the
TCA and the ETC. Eight NADH binding sites exist in the aerobic metabolic
pathways. Enzymes for NADH binding are: (1) glyceraldehyde- 3-phosphate
dehydrogenase; (2) lactate dehydrogenase; (3) pyruvate dehydrogenase complex; (4) isocitrate dehydrogenase; (5) α-ketoglutarate dehydrogenase complex; (6) malate dehydrogenase; (7) complex I of the electron transport chain;
(8) cytosolic malate dehydrogenase; and (9) cytosolic glycerol-3-phosphate
dehydrogenase.
I. INTRODUCTION
ICOTINAMIDE adenine dinucleotide (NAD/NADH) is
an important coenzyme involved in the energy metabolism
of living cells. NADH binds to proteins associated with both
aerobic and anaerobic metabolism, as shown in Fig. 1. In
anaerobic metabolism (glucose to lactate), two protein-binding
sites are present for NADH binding [1]. In contrast, in aerobic metabolism (glucose to H2 O and CO2 ), eight proteinbinding sites are present for NADH binding [1]. The ratio of
free to protein-bound NADH reflects different metabolic states
N
Manuscript received September 5, 2007; revised October 9, 2007. This work
was supported in part by the Whitaker Foundation under Grant RG-99-0127 and
in part by the Skin Cancer Foundation under Grant 582019.
Y. Yuan was with the Department of Bioengineering, The University of
Toledo, Toledo, OH 43606 USA. He is now with the Department of Engineering Physics, McMaster University, Hamilton, ON L8 S 4L7, Canada (e-mail:
yuanye@mcmaster.ca).
Y. Li was with the Department of Bioengineering, The University of Toledo,
Toledo, OH 43606 USA (e-mail: yfll@hotmail.com).
B. D. Cameron and P. Relue are with the Department of Bioengineering, The University of Toledo, Toledo, OH 43606 USA (e-mail:
bcameron@eng.utoledo.edu; prelue@eng.utoledo.edu).
Color versions of one or more of the figures in this paper are available online
at http://ieeexplore.ieee.org.
Digital Object Identifier 10.1109/JSTQE.2007.910806
in cells associated with different levels of aerobic and anaerobic
metabolism [2]–[4].
Glycolysis and respiration are tightly coupled in normal
cells. However, cancer cells lack the ability to integrate energy metabolism between glycolysis and respiration. Metabolic
changes of cancer cells include enhanced glycolysis, excessive
lactate production, and reduced respiratory capacity [2], [5]–[8].
Rapidly growing cancer cells can derive about 60% of their energy from glycolysis and 40% of their energy from oxidative
phosphorylation [9], whereas normal cells derive less than 10%
of their energy from glycolysis and more than 90% of their
energy from oxidative phosphorylation [10]. The shift in the energy metabolism of cancer cells from respiration to aerobic glycolysis may reduce the interaction of NADH with NAD-linked
dehydrogenases, leading to a higher ratio of free to proteinbound NADH in the cancer cells. Further, the NAD-linked dehydrogenases involved in glycolysis are free in the cytosol [1],
and most of the NAD-linked dehydrogenases involved in respiration are located in mitochondria, and two of them, namely,
pyruvate dehydrogenase and the complex I, are attached to the
inner mitochondrial membrane [1]. The high viscosity of the
mitochondrial matrix space [11] and the association of citric
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IEEE JOURNAL OF SELECTED TOPICS IN QUANTUM ELECTRONICS, VOL. 13, NO. 6, NOVEMBER/DECEMBER 2007
acid cycle (TCA) and electron transport chain (ETC) enzymes
with one another [12], [13] and with the inner mitochondrial
membranes [11] significantly reduce the mobility of NADH;
thus, the metabolic shift in cancer cells may provide a much
more relaxed environment for NADH motion.
NADH is a natural fluorophore that emits in the blue end of
the visible light spectrum upon ultraviolet (UV) light excitation.
The fluorescence properties of cellular NADH were first studied
by Chance and coworkers as an optimal indicator of intracellular oxidation–reduction states and a useful tool to study mitochondrial function in energy-linked processes [14], [15]. Since
Chance’s pioneering work, cellular NADH fluorescence has
been widely studied for its ability to probe cell metabolic conditions [4], [16], proliferation [17], differentiation [18], transformation [2], [3], [19], [20], and drug response [21], [22].
The properties of the free and protein-bound NADH in living cells have been studied by the methods of fluorescence
spectral analysis [19], [23], [24], fluorescence lifetime measurements [16], [20], [23], [25]–[28], and time-resolved fluorescence
anisotropy decay [29]. When bound to protein, the wavelength of
maximum NADH emission is shifted from 460 to 440 nm [30].
This emission property has been applied to study the ratio of
free to protein-bound NADH in cells through spectral fitting
analysis of the fluorescence emission of the cells [2]–[4]. When
NADH binds to protein, the fluorescence lifetime is generally
increased [25], [26], [29]. However, the fluorescence lifetime of
protein-bound NADH varies depending on the protein to which
the NADH binds [27], [29]. Some NADH/protein complexes, in
fact, have fluorescence lifetimes comparable to or even shorter
than the free NADH [28], [29], thus, increasing the challenge
of using fluorescence lifetime measurements to discriminate
between free and protein-bound NADH. Time-resolved fluorescence anisotropy decay has been used to address this problem
since anisotropy-based methods target the rotational mobility of
a fluorophore [29]. The large difference in rotational diffusion
resulting from the difference in size between the free NADH and
the NADH/protein complexes provides an excellent contrast for
time-resolved fluorescence anisotropy measurements, allowing
successful resolution of free and protein-bound NADH sharing
comparable lifetimes [29].
In solution, rotational diffusion is a dominant cause of fluorescence depolarization [31]. Thus, fluorescence anisotropy,
which is a measure of retained polarization, increases when a
fluorophore associates with a larger molecule or is observed in a
solution with higher viscosity [31]. However, in cells and tissue,
both scattering and absorption can significantly contribute to the
depolarization of both excitation and emission light, and hence,
the measured fluorescence anisotropy.
To test the feasibility of using fluorescence anisotropy
measurements to detect changes in NADH during metabolic
perturbation, rotenone was employed to block mitochondriaassociated energy metabolism in cells. Rotenone blocks electron transport and induces NADH accumulation in mitochondria, which in turn, increases NADH fluorescence intensity [2],
[19], [26]. Fluorescence lifetime measurements have shown that
rotenone-treated endothelial cells have a decreased fluorescence
lifetime as compared to control cells, suggesting an accumula-
Fig. 2. Schematic diagram of the experimental setup for fluorescence and
reflectance spectroscopy. For fluorescence spectroscopy, fiber 1 was used for
illumination. For reflectance spectroscopy, fiber 3 was used for illumination.
Fiber 2 was used to deliver either fluorescence emission or reflectance from the
sample to the spectrometer.
tion of free NADH within the mitochondria [26]. Therefore,
rotenone-treated cells are expected to have higher fluorescence
emission and lower fluorescence anisotropy as compared to control cells.
An additional component of this study was to determine if
differences in metabolic properties between normal human skin
melanocytes and melanoma cells could be detected with fluorescence anisotropy measurements. The fluorescence signal from
these cell types is very different due to differences in scattering, absorption, the total quantity, and the state of NADH (free
or protein bound). Although the fluorescence anisotropy is independent of the total quantity of NADH, it is dependent on
the state of NADH as well as by both scattering and absorption behaviors. Absorption and scattering properties of the cell
suspensions were studied using reflectance spectroscopy. After
correction for variation in scattering and absorption, fluorescence anisotropy between cell types was compared.
II. MATERIALS AND METHODS
A. Experimental Setup
The experimental setup for fluorescence and reflectance spectroscopy is shown in Fig. 2. A 100 W mercury arc lamp
(Osram, Danvers, MA) was used as a light source. Excitation
light was provided using a 360 nm excitation bandpass filter
with a 20 nm bandwidth (Chroma Technology, Rockingham,
VT). A 390-nm dichroic mirror was used to reflect the excitation light. Two silica-core optical fibers (Thorlabs, Newton, NJ)
with diameter of 1 mm were used to deliver excitation light and
collect fluorescence emissions. The numerical aperture of the
fibers was 0.22 ± 0.02. The distance of the fiber tip to the sample surface was 1 cm for both the illumination and collection
fibers. The angle between the illumination and collection fibers
was 45◦ . The polarizer was a UV passing glass linear polarizer
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YUAN et al.: FLUORESCENCE ANISOTROPY OF CELLULAR NADH
(Edmund Optics, Barrington, NJ). The analyzer consisted of a
pair of film linear polarizers (Edmund Optics, Barrington, NJ)
oriented orthogonally to one another. The film polarizers transmitted wavelengths greater than 380 nm. A custom-designed
holder was used for the two fibers, the polarizer and the analyzer
to ensure reproducible positioning. The collection fiber was connected to a USB2000 fiber optic spectrometer (Ocean Optics,
Dunedin, FL) as the detector. Both the filter wheel and the spectrometer were controlled by a PC. Fluorescence and reflectance
spectra collected by the spectrometer were displayed and stored
on the PC. To collect non-polarized spectra, the linear polarizers were removed from the light path. For reflectance spectroscopy, a 6.5 W LS-1 tungsten halogen light source (Ocean
Optics, Dunedin, FL) with a 1.0 neutral density filter was used.
A silica-core optical fiber with diameter of 1 mm was used to
deliver the halogen light.
B. Cell Culture
Human melanocyte cell strain HEMn_LP (Cascade Biologics, Portland, OR) was cultured with Medium 154 (Cascade Biologics, Portland, OR). Human melanoma cell lines WM793
and WM115 (The Wistar Institute, Philadelphia, PA) were
cultured with Dulbecco’s modified Eagle’s medium (DMEM,
ICN Biomedical Inc, Aurora, OH) supplemented with 10% fetal bovine serum (Gibco) and 5 µg/mL insulin (Sigma). Both
WM793 and WM115 are vertical growth phase melanoma. All
of the cells were routinely cultured in a 37 ◦ C humidified 5%
CO2 incubator (NAPCO).
For spectra collection, cultured cells were trypsinized when
they reached near or 100% confluence. The cells were washed
three times with phosphate buffered saline (PBS). The third
wash was measured to account for background fluorescence.
Cell concentrations were measured with a hemocytometer and
cell viabilities were determined by trypan blue exclusion assay.
C. Rotenone Treatment
The cells were resuspended in their culture media after
trypsinization. The cell concentrations of the suspensions were
8 × 106 to 1 × 107 cells/mL for the melanocytes, 8 × 106
cells/mL for WM115, and 1 × 107 cells/mL for WM793. A
1 mL aliquot of cell suspension was split evenly into two sterile 1.5 mL centrifuge tubes. Cells were allowed to settle for
2–3 min, and 2.5 µL of supernatant was removed from each
tube. A 2.5 µL aliquot of 1 mM rotenone (Sigma-Aldrich) stock
solution was added and mixed with the cell suspension in one
tube. As the rotenone stock solution was made by dissolving
rotenone in ethanol, 2.5 µL of ethanol was added and mixed
with the cell suspension in the other tube as a control. The cell
suspensions were incubated for 10 min at room temperature and
gently swirled to prevent cell precipitation. Following the incubation, the cells were spun down, washed three times with PBS,
and finally resuspended in 0.5 mL PBS in each tube for fluorescence measurements. Cell viabilities were checked before the
rotenone treatment and after the fluorescence measurements.
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D. Fluorescence and Reflectance Spectroscopy of Cell
Suspension
Due to the different scattering and absorption properties of
the cell suspensions, the melanocytes and melanoma cells were
resuspended in the PBS at different concentrations ranging from
5 × 106 to 6 × 107 cells/mL. For the spectral measurement of
each cell suspension, 0.2 mL of cell suspension was transferred
into a 0.2 mL custom-made mini petri dish. After waiting for
5 min to allow the cells to settle, sequential polarized fluorescence spectra were collected with the polarization orientation
between the polarizer and analyzer set as parallel, perpendicular,
perpendicular, and parallel. The two parallel polarized fluorescence spectra were compared to determine if photobleaching
occurred. Polarized reflectance spectra were collected in the
same manner immediately following the collection of the polarized fluorescence spectra. Nonpolarized reflectance spectra
were collected by removing the polarizers from the light path.
The nonpolarized reflectance spectra were used to study the
changes of absorption of the cell suspensions of different cell
types at different concentrations. Cell viabilities were checked
before and after all of the fluorescence and reflectance spectra
were collected.
Fluorescence and reflectance spectra were imported into
MATLAB. Polarized spectra for reflectance and fluorescence
were calculated by averaging the two parallel or two perpendicular polarized spectra collected in each experiment. The fluorescence anisotropy (FA) and degree of linear polarization (DLP)
were calculated as
FA = (If // − If ⊥ )/(If // + 2If ⊥ )
(1)
DLP = (Ir // − Ir ⊥ )/(Ir // + Ir ⊥ )
(2)
where I is the intensity of a spectrum, and f or r refer to fluorescence or reflectance spectra, respectively, and // or ⊥ represent
either parallel or perpendicular alignment of the polarizer and
analyzer [31].
For experiments with rotenone treatment, the total fluorescence spectrum, If (λ), was calculated from the parallel and
perpendicular spectra as
If (λ) = If // (λ) + 2If ⊥ (λ).
(3)
The total fluorescence was calculated by integrating the total
fluorescence spectrum If (λ) over the wavelength range from
430 to 550 nm. The change in the fluorescence due to rotenone
treatment was calculated as the difference between the total
fluorescence of the rotenone-treated cells and the control cells
divided by the total fluorescence of the control cells.
III. RESULTS
A. Effects of Rotenone Treatment on Cell Fluorescence
Fluorescence spectra with polarized excitation at 360 nm
were collected from melanocyte, WM115, and WM793 cell
suspensions on different days. No background fluorescence
was detected and no photobleaching was observed during the
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TABLE I
CELL VIABILITIES BEFORE ROTENONE TREATMENT AND AFTER
FLUORESCENCE MEASUREMENT
Fig. 4. Reflectance spectra of melanocyte (left), WM115 (middle), and
WM793 (right) cell suspensions at different cell concentrations.
TABLE II
CHANGE IN TOTAL FLUORESCENCE FOR THE DIFFERENT CELL TYPES WITH
AND WITHOUT ROTENONE TREATMENT
Fig. 3. Fluorescence anisotropy of melanocyte (left), WM115 (middle), and
WM793 (right) cell suspensions. Fluorescence anisotropies decreased with
rotenone treatment for all three cell types.
experiments. The cell viabilities for the three cell types were always greater than 90% (Table I). After rotenone treatment, cell
fluorescence from all cell types increased appreciably relative to
the control cells. The change in total fluorescence with rotenone
treatment was calculated as described in Section II. As shown in
Table II, the average increase in total fluorescence ranged from
35.8 to 43.8% for the three cell types; no significant difference
is seen between cell types.
The fluorescence anisotropies were calculated using (1) as
described in Section II for cells with and without rotenone
treatment. The fluorescence anisotropy decreased with rotenone
treatment in all three cell types tested as shown in Fig. 3.
B. Reflectance Spectra of the Cell Suspensions
Differences in pigmentation levels between the melanocytes
and melanoma cells used in this study resulted in a large difference in absorption between the cells. To examine these differences, nonpolarized reflectance spectra were collected from the
melanocyte and melanoma cell suspensions at concentrations
ranging from 5 × 106 to 6 × 107 cells/mL. The reflectance spectra collected for each cell type and concentration were averaged
over the measurements of different batches of cells collected
on different days. Each average spectrum was then normalized
against its peak intensity. The normalized reflectance spectra are
shown in Fig. 4. All three cell types show stronger absorption
at higher cell concentrations, which results in a red shift of the
reflectance spectra. The red shift was much more pronounced
for the melanocytes than for the two melanoma cell lines.
The peak wavelength was also determined for each spectrum and averaged over all spectra collected for each cell type
and concentration. The values of peak wavelength are summarized in Table III. The peak wavelength was red shifted with
increasing concentration of the cell suspensions for each cell
type. All three cell types showed a significant red shift of
the peak wavelength as the cell concentration increased from
2 × 107 to 6 × 107 cells/mL (P < 0.05). The melanocytes also
showed a significant red shift between 1 × 107 and 2 × 107
cells/mL (P < 0.05). However, no statistically significant shift
in peak wavelength was observed for the cell concentrations
from 5 × 106 to 1 × 107 cells/mL. The lack of red shift at the
low cell concentrations suggests that absorption is considerably
reduced.
No statistical difference was seen between the peak wavelengths of WM115 and WM793 at any concentration, suggesting similar absorption by these two melanoma cell lines.
However, the peak wavelength of the melanocytes was significantly higher than either of the two melanoma cell lines at
the cell concentrations of 6 × 107 (P < 0.0001) and 2 × 107
cells/mL (P < 0.05), suggesting a much stronger absorption
by the melanocytes than the melanoma cells at these high cell
concentrations. There was no statistically significant difference
in the peak wavelength between the melanocytes and melanoma
cells at the concentrations of 1 × 107 and 5 × 106 cells/mL,
suggesting that the melanocytes and melanoma cells shared a
similar absorption at these low cell concentrations.
C. Polarized Reflectance Spectra of the Cell Suspensions
To determine scattering properties of the three cell types as
a function of concentration, polarized reflectance spectra were
collected immediately after collection of the polarized fluorescence spectra. The DLP was calculated using (2) as described
in the Section II. The DLP provides information about multiple scattering within a turbid media, where increased scattering
tends to depolarize incident polarized light to a greater extent,
leading to a lower DLP. The DLP was averaged over different
batches of cells of the same type and concentration collected on
different days. As shown in Fig. 5, the DLP decreased as the
cell concentration increased for the three cell types, indicating
that as the cell layer increased in thickness, depolarization of
the backscattered light increased.
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YUAN et al.: FLUORESCENCE ANISOTROPY OF CELLULAR NADH
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TABLE III
EFFECT OF CELL CONCENTRATION ON THE PEAK WAVELENGTH OF THE REFLECTANCE SPECTRA OF THE CELL SUSPENSIONS
Fig. 5. DLP of the melanocyte (left), WM115 (middle), and WM793 (right)
cell suspensions with different cell concentrations. Average DLP was calculated
from the measurements on different batches of cells. The cell concentrations
(cells/mL) and numbers of batches of cells are shown in the legends.
As shown in Fig. 6(a) and (b), the DLP of the melanocytes
was higher than that of the two melanoma cell lines at the
two highest concentrations. These are the same concentrations
where the melanocytes showed significantly stronger absorption
than the melanoma cells. Similar results were reported in the
literature [32], [33], where the authors suggest that stronger absorption results in a higher DLP due to the absorption of highly
scattered photons with long optical pathlengths. At low cell concentrations (< 1 × 107 cells/mL), absorption by the cells was
relatively weak, and the absorption of the melanocytes and the
two melanoma cells was not statistically different (see Table III).
When comparing the cells with similar absorption, such as the
WM115 and WM793 cells at equal concentration, the DLP is
not the same. This suggests that the difference in DLP is not
associated with absorption, but with differences in scattering.
Therefore, the difference in DLP seen between the melanocytes
and melanoma cells when their absorptions are equivalent cannot be attributed to absorption.
The DLP of WM115 cells was always lower than that of
WM793 cells at the same concentrations, suggesting a stronger
scattering of WM115 cells than WM793 cells. At the highest
cell concentrations where absorption was significant, the DLP
of the melanocytes was higher than that of the two melanoma
cell lines [Fig. 6(a) and (b)]. However, as the concentration
was reduced to 1 × 107 cells/mL, the DLP of the melanocytes
[Fig. 6(c)] was between that of the two melanoma cell lines,
suggesting that absorption no longer played a significant role in
the DLP. The DLP of the melanocytes at the concentration of
1 × 107 cells/mL was practically the same as that of WM115
Fig. 6. Comparison of the DLP between different cell types at the same cell
concentrations (a)–(c). DLP spectra were averaged over the measurements of
different batches of cells. (a) DLP for the cell suspensions of 6 × 107 cells/mL.
(b) DLP for the cell suspensions of 2 × 107 cells/mL. (c) DLP for the cell
suspensions of 1 × 10 7 cells/mL. (d) DLP for the cell suspensions of 1 ×
10 7 cells/mL for the melanocytes and WM793 cells and 7 × 106 cells/mL for
WM115 cells. Dashed lines represent ±1 standard deviation for the dataset.
cells at 7 × 106 cells/mL [see Fig. 6(d)], suggesting that the
same scattering property could be obtained by lowering the cell
concentration for WM115 cells.
D. Polarized Fluorescence Spectra of the Cell Suspensions
Polarized fluorescence spectra were collected from the three
cell types at different cell concentrations. Cell viabilities were
checked before and after each measurement and were always
greater than 90%. Fluorescence anisotropy was calculated using
(1) as described in the Section II, and the FA was averaged over
different batches of cells for each cell type and concentration.
As shown in Fig. 7(a)–(c), fluorescence anisotropy decreased
as the cell concentration increased for the three cell types, suggesting a dependence of fluorescence anisotropy on the scattering of the cell suspensions. The fluorescence anisotropy,
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Fig. 8. Fluorescence ratio spectra of the different cell types. Fluorescence
ratio spectra were averaged over six batches of cells for each of the three cell
types, and normalized against the maximum intensities.
TABLE IV
FLUORESCENCE INTENSITY RATIO AT 430 AND 465 NM FOR THE DIFFERENT
CELL TYPES
Fig. 7. Comparison of the fluorescence anisotropy of the cell suspensions with
different cell concentrations (a)–(c) and between different cell types (d). Fluorescence anisotropies were averaged over measurements of different batches of
cells. (a) Fluorescence anisotropy of melanocyte suspensions with concentration from 1 × 10 7 to 6 × 10 7 cells/mL. (b) Fluorescence anisotropy of WM115
cell suspensions with concentration from 7 × 106 to 6 × 10 7 cells/mL. (c)
Fluorescence anisotropy of WM793 cell suspensions with concentration from
1 × 10 7 to 6 × 10 7 cells/mL. (d) Comparison of fluorescence anisotropy between the melanocytes (1 × 10 7 cells/mL), WM115 cells (7 × 10 6 cells/mL),
and WM793 cells (1 × 10 7 cells/mL). Dashed lines represent ±1 standard deviation for the dataset.
like the DLP, was remarkably higher for the melanocytes
than for the two melanoma cell lines at high cell concentrations (> 2 × 107 cell/mL). This difference could partly be due
to the much stronger absorption of the melanocytes relative
to the melanoma cells at the high cell concentrations. However, the fluorescence anisotropy of the melanocytes was still
appreciably higher than that of the two melanoma cell lines
at the lowest cell concentrations [Fig. 7(d)], where the two
melanoma cell lines showed similar fluorescence anisotropy.
The DLP measurements [Fig. 6(d)] showed that all three cell
types had similar scattering properties at these cell concentrations. Therefore, the higher fluorescence anisotropy for the
melanocytes is not due to differences in scattering and/or absorption of the cell suspensions. As a result, the higher fluorescence
anisotropy for the melanocytes could indicate a higher proportion of protein-bound NADH and/or a more stringent microenvironment for NADH in the melanocytes than in the melanoma
cells.
E. Fluorescence Intensity Ratio of Melanocytes and Melanoma
Cells
Fluorescence emission of the protein-bound NADH is blue
shifted as compared to that of the free NADH. It was suggested
that the ratio of fluorescence intensity at 410 and 465 nm could
provide a sensitive test of NADH binding [19]. Due to an artifact in our data at 410 nm that was caused by the mini petri
dish, we instead compare the ratio of fluorescence intensity at
430 and 465 nm. Cell concentrations were 1 × 107 cells/mL for
the melanocytes and WM793 cells and 7 × 106 cells/mL for
WM115 cells. Polarized fluorescence spectra were defined as
the difference between the parallel and perpendicular fluorescence spectra. Polarized reflectance spectra were defined as the
difference between the parallel and perpendicular reflectance
spectra. Fluorescence ratio spectra were obtained by normalizing the polarized fluorescence spectra with its corresponding
polarized reflectance spectra. This normalization was experimentally shown to recover intrinsic fluorescence of fluorophores
from turbid media [34]. As shown in Fig. 8, fluorescence ratio
spectra were averaged over six batches of cells for each of the
three cell types and normalized against the maximum intensities. The emission peaks were around 420 nm. However, the
fluorescence ratio spectrum of the melanocytes showed an appreciable blue shift as compared to those of the two melanoma
cell lines.
The ratios of fluorescence intensity at 430 and 465 nm for
the melanocytes and melanoma cells are shown in Table IV.
The fluorescence intensity at each wavelength (i.e., 430 or 465
nm) was obtained by integrating the fluorescence ratio spectra over a wavelength range of ±1 nm about the wavelength.
The fluorescence intensity ratio was averaged over six batches
of cells for each cell type. The melanocytes demonstrated a
significantly higher fluorescence intensity ratio than the two
melanoma cell lines (P < 0.05), while no statistical difference
was seen between the two melanoma cell lines. The higher
fluorescence intensity ratio of the melanocytes suggests that
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YUAN et al.: FLUORESCENCE ANISOTROPY OF CELLULAR NADH
the NADH binding was higher in the melanocytes than in the
melanoma cells, which is consistent with the higher fluorescence
anisotropy of the melanocytes as compared to the melanoma
cells.
IV. DISCUSSION
To evaluate the feasibility of using fluorescence anisotropy of
cellular NADH for tracking metabolic changes in living cells,
rotenone treatment was applied to induce metabolic changes
in the cells. Rotenone inhibits the complex I of the electron
transport chain [35], and thus, blocks the transfer of electrons
through the electron transport chain, which in turn, arrests the
oxidation of NADH at the site of the complex I. Therefore,
NADH accumulates in the mitochondria with rotenone treatment. Studies have demonstrated an increase of NADH fluorescence emission after rotenone inhibition in various types of
cells [2], [19], [26]. In addition, spectral fitting analysis of the
fluorescence of rotenone-treated fibroblasts has revealed an increased free/bound NADH ratio as compared to the untreated
cells [2]. Fluorescence lifetime imaging has also demonstrated a
decreased fluorescence lifetime for rotenone-treated endothelial
cells compared to control cells, which was attributed to the accumulation of free NADH within the mitochondria [26]. A recent
study has shown that free NADH constitutes about 60% of the total matrix NADH in intact mitochondria, and the rest of NADH is
protein-bound and contributes to about 80% of the fluorescence
emission [27]. As the total matrix NADH level increases, the
pool of the protein-bound NADH saturates, whereas the pool of
free NADH increases linearly as the total NADH level increases
[27]. The mitochondrial NADH level increases with rotenone inhibition. Thus, the protein-bound NADH pool in the mitochondria could saturate, while the free NADH pool in the mitochondria could still linearly increase. Therefore, in our experiments
with rotenone treatment, the increase in overall fluorescence
could be attributed to an increase in both free and protein-bound
NADH, whereas a relative increase in the free NADH pool could
account for the decreased fluorescence anisotropy seen in all cell
types.
Based on the assumption that enhanced glycolysis and reduced aerobic respiration of cancer cells may change the physicochemical properties of NADH, fluorescence anisotropy of the
cellular NADH of the melanocytes and melanoma cells were
collected to probe the different metabolic properties of the
cells. In order to compare fluorescence anisotropy between
the different types of cells, differences in optical properties of
the cell suspensions, namely, absorption and scattering, need
to be addressed. Reflectance spectroscopy was used to compare the optical properties of the melanocyte and melanoma
cell suspensions. The melanocytes showed much stronger absorption than the melanoma cells at high cell concentrations
(> 2 × 107 cell/mL), resulting in a higher degree of linear polarization and higher fluorescence anisotropy for the melanocytes.
The effect of absorption on the DLP and fluorescence anisotropy
may be caused by the loss of highly scattered long pathlength
photons in the backscattered light [32], [33]. The difference
in absorption between the melanocytes and the melanoma
cells was minimized with dilution of the cell suspensions,
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and the relatively weak absorption at the lowest cell concentrations might be eliminated through normalization. WM115
cells showed the strongest scattering and WM793 cells showed
the weakest scattering among the three cell types. The different scattering properties may be caused by the different
level and shape of the index mismatched structures in these
cells.
When the cell concentrations were adjusted to minimize differences in scattering and absorption effects, the melanocytes
showed an appreciably higher fluorescence anisotropy than
either of the melanoma cell lines. Thus, this higher fluorescence anisotropy suggests a higher proportion of protein-bound
NADH and/or a more stringent microenvironment for NADH
in the melanocytes. The ratio of fluorescence intensity at 430
and 465 nm also supports higher levels of NADH binding in the
melanocytes. As described in the Section I, two NADH binding sites exist for anaerobic metabolism, while eight NADH
binding sites exist for aerobic metabolism. The mitochondrial matrix also provides a more mobility-restricted environment than the cytosol for proteins or even small molecules
like NADH [11]. Thus, the metabolic shift of normal aerobic metabolism to enhanced glycolysis and reduced respiration may decrease the binding of NADH to metabolic proteins, and provide a more mobile microenvironment for the
NADH.
Aerobic glycolysis, which is a characteristic of many tumors,
is accompanied by significantly enhanced activities of glycolytic
enzymes and reduced activities of the enzymes of the TCA cycle and the electron transport chain [8], [36]–[38]. Melanoma
cells have shown the elevated expression of c-myc oncogene in
comparison to melanocytes [39]. The activation of the c-myc
oncogene results in an oncogenic transcription factor (c-Myc),
which is capable of activating gene expressions for the glucose transporter, glycolytic enzymes, and the lactate dehydrogenase [40]–[42]. Decreased enzymes of the electron transport
chain have also been observed in melanoma [43]. Therefore, the
enhanced glycolysis and reduced respiration of melanoma cells
may result in a lower proportion of NADH binding to proteins
and a more mobile microenvironment for NADH, resulting in
a decrease in fluorescence anisotropy as observed in this study.
Several other studies on cellular NADH fluorescence have revealed reduced binding of NADH in cancer cells relative to their
normal counterparts [2], [19], [28].
Since aerobic glycolysis is constitutively upregulated in
tumors, cells derived from tumors typically maintain their
metabolic phenotypes in culture under normoxic conditions [7].
Thus, the in vitro fluorescence anisotropy measurements of cellular NADH in cultured cells may mirror behavior of in vivo
tissues, rendering the possibility of extending the fluorescence
anisotropy method to in vivo imaging. Hypoxia in tumor tissues induces anaerobic glycolysis, which will further increase
the contrast for the fluorescence anisotropy measurements. Progressively increased glycolytic rate and reduced respiration capacity are associated with cancer progression [44]–[46]. Therefore, fluorescence anisotropy measurement of cellular NADH
may potentially allow discrimination of cancer cells at different
stages of progression.
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IEEE JOURNAL OF SELECTED TOPICS IN QUANTUM ELECTRONICS, VOL. 13, NO. 6, NOVEMBER/DECEMBER 2007
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YUAN et al.: FLUORESCENCE ANISOTROPY OF CELLULAR NADH
Ye Yuan was born in Leshan City, Sichuan Province, China. He received the
B.S. degree in biochemical engineering from Chengdu University of Science
and Technology, Chengdu, China, in 1992, the M.S. degree in fermentation
engineering from Wuxi University of Light Industry, Wuxi, China, in 1999, and
the Ph.D. degree in bioengineering from the University of Toledo, Toledo, OH,
in 2006.
He was engaged in biopharmaceutical industry in China. He is currently a
Postdoctoral Researcher in the Biophotonics Laboratory, McMaster University,
ON, Canada. His current research interests include general field of biophotonics.
1679
Patricia Relue received the Ph.D. degree in chemical engineering from the
University of Michigan, Ann Arbor, in 1994.
In 1993, she joined the University of Toledo, Toledo, OH, as an Assistant Professor of chemical engineering, where she is currently working as an Associate
Professor with the Graduate Program Director of bioengineering. Her current
research interests include cellular engineering, optical imaging, and bioethanol
production.
Dr. Relue was the recipient of the University and College Outstanding Teaching Awards as well as teaching awards from the Biomedical Engineering Society
Student Chapter.
Yanfang Li received the B.S. and M.S. degrees in biomedical engineering from
Tianjin University, Tianjin, China, in 1997 and 2000, respectively, and the Ph.D.
degree in bioengineering from the University of Toledo, Toledo, OH, in 2005.
Brent D. Cameron was born in Sioux City, IA, on December 29, 1970. He received the B.S., M.S., and Ph.D. degrees in biomedical engineering from Texas
A&M University, College Station, in 1994, 1996, and 2000, respectively.
From August 2000 to 2005, he was an Assistant Professor in the Department of Bioengineering, the University of Toledo, Toledo, OH. In 2006, he was
promoted to the rank of Associate Professor with tenure, where he founded the
Biomedical Optics Laboratory in the Department of Bioengineering. His current
research interests include the use of optical methods for medical diagnostics and
sensing including theoretical modeling and also specialized in optical polarization, fluorescence sensing, Raman spectroscopy, and Monte Carlo modeling of
photon migration.
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