Reduced Nicotinamide Adenine Dinucleotide (NADH)

Anti-Cancer Agents in Medicinal Chemistry, 2009, 9, 000-000

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Reduced Nicotinamide Adenine Dinucleotide (NADH) Fluorescence for the Detection of Cell Death Hsing-Wen Wang1,*, Yau-Huei Wei2,3 and Han-Wen Guo1 1 3

Institute of Biophotonics, 2Institute of Biochemistry and Molecular Biology, National Yang-Ming University, Taipei, Taiwan; Department of Medicine, Mackay Medical College, Tanshui, Taipei County, Taiwan Abstract: NADH/FAD fluorescence spectroscopy/imaging is an extremely useful tool to probe cellular metabolism and has been applied in the clinic such as early cancer detection. Recently, the potential of using NADH/FAD fluorescence as a biomarker to detect cell death has been investigated for development of cancer treatments with higher efficacy. This review aims to provide the updated information in cell death detection using the NADH/FAD fluorescence spectroscopy and imaging based on measurement of the intensity or lifetime of NADH or FAD fluorescence. The response of NADH fluorescence lifetime to metabolic perturbation, hypoxic environment, and anaerobic glycolysis (e.g., in precancerous tissues and stem cells) is also reviewed to discuss the nature and implications of the lifetime change of NADH fluorescence. Further studies are required to understand the actual site and mechanism of NADH binding of a specific death pathway for future successful in vivo detection of cell death using the NADH fluorescence lifetime.

Keywords: NAD(P)H and FAD fluorescence intensity and lifetime, redox ratio, mitochondrial respiration, cell death. INTRODUCTION The intrinsic fluorescence of biological molecules includes reduced nicotinamide adenine dinucleotide (NADH), flavin adenine dinucleotide (FAD), structural proteins such as collagen, elastin and their cross links, the aromatic amino acids including tryptophan, tyrosine and phenylalanine, and porphyrins. Among these intrinsic fluorophores, the NADH and FAD as the principal coenzymes in many metabolic enzymes make them a convenient in vivo and noninvasive fluorescent probe of the tissue metabolic state as pioneered by Chance and colleagues since 1962 [1]. In the past decade, the intrinsic fluorescence of NADH and FAD has been substantially used as one of the early diagnostic tools for colon, cervical, esophageal, lung, bladder, and breast cancers [2]. The underlying mechanism is that neoplastic cells have an increased metabolic demand due to rapid cell division compared with normal cells [3, 4]. Another mechanism, also the hallmark of cancers, involves the so-called Warburg effect that cellular energy supply shifts from a much more efficient aerobic metabolism to a less efficient anaerobic glycolysis [5]. These metabolic changes would reflect changes in the relative concentrations of NADH and FAD as normal tissues progress to cancers, and thus can be monitored noninvasively and without exogenous labels using optical techniques. The Warburg effect was observed during differentiation of stem cells in that stem cells are similar to cancer cells in terms of the use of glycolysis rather than aerobic metabolism for the major supply of energy [6]. A previous study has demonstrated that NADH together with FAD fluorescence can be probably used as a noninvasive biomarker for the detection of stem cells [7]. Mitochondria participate in many important biochemical reactions. Structurally, the organelles have four compartments: the outer membrane, the inner membrane, the intermembrane space, and the matrix. Their principal biological function of mitochondria is energy transduction and they are called the powerhouse of mammalian cells because a majority of adenosine 5-triphosphate (ATP) is generated by means of respiration and oxidative phosphorylation, in which a proton gradient is generated across the inner membrane of mitochondria by respiratory enzyme Complexes I, III, and IV. This proton gradient is used to drive ATP synthesis by the phosphorylation of adenosine 5’-diphosphate (ADP) through Complex V (ATP synthase). NADH and reduced flavin adenine dinucleotide *Address correspondence to this author at the Institute of Biophotonics, National Yang-Ming University, 155 Li-Nong St., Sec. 2, Taipei 11221, Taiwan; Tel: 886-2-2826-7378; Fax: 886-2-2823-5460; E-mail: [email protected] 1871-5206/09 $55.00+.00

(FADH2) are the principal electron donors during this process. High energy electrons (or hydrogen ions derived from NADH and FADH2) are transported along the respiratory enzyme complexes to molecular oxygen to produce water and ATP, and concurrently NADH and FADH2 are oxidized in Complex I (NADH:coenzyme Q oxidoreductase) and Complex II (succinate: coenzyme Q oxidoreductase), respectively. Thus, generation of ATP in the mitochondrial inner membrane is coupled to the oxidation of NADH and FADH2. NAD+ is also a coenzyme required by glyceraldehyde 3-phosphate dehydrogenase in glycolysis that occurs in the cytosol. In glycolysis, a mole of glucose is metabolized to two moles of pyruvate and two moles each of NADH and ATP are generated. Under aerobic conditions, pyruvate continues to enter the mitochondria for more efficient production of ATP. NADH generated in glycolysis can enter mitochondria by aspartate-malate shuttle or glycerol phosphate shuttle, and is then oxidized via the respiratory chain to generate additional ATP. The reduced pyridine nucleotides (NADH and NADPH) and oxidized FAD emit fluorescence, but their oxidized (NAD+) and reduced (FADH2) form, respectively, do not. The NAD(P)H and FAD have excitation maxima at ~350 and 450 and emission maxima at ~450 and 535 nm, respectively, when fluorescence excitation-emission matrices (EEMs) are performed [8, 9]. A fluorescence EEM is a two dimensional contour plot, which displays the fluorescence intensities in a range of excitation and emission wavelengths so that both optimal excitation and emission wavelengths can be obtained for use in spectrofluorometric or fluorescence microscopic measurement. Both NADH and NADPH have nondiscriminative spectral features. However, NADH dominates cellular fluorescence over NADPH because studies have shown that in mammalian cells the concentration of NADH is ~5 times greater than that of NADPH and the quantum yield of mitochondrial NADH is 1.25 to 2.5 times higher than that of NADPH [10]. NADPH also represents a minor and roughly constant fluorescence background with respect to metabolic perturbations that only a minor (~10-15%) contribution of fluorescence change from NADPH during the normoxic- anoxic transition in tissue analyses of pyridine nucleotide content in heart and liver [11, 12]. Thus, NADH dominates the fluorescence changes due to metabolic perturbations. In addition, it is thought that the fluorescence of NAD(P)H comes mainly from mitochondria in mammalian cells. Blinova et al. [13, 14] demonstrated that this dominated NADH fluorescence intensity is specifically attributed to those NADH bound to Complex I proteins. The mitochondrial function in vivo evaluated by NADH fluo-

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2 Anti-Cancer Agents in Medicinal Chemistry, 2009, Vol. 9, No. 9

rescence intensity has recently been reviewed by Mayevsky and Rogatsky [15]. Because fluorescence intensity measurement may be affected by instrumentation parameters and attenuated by hemoglobin absorption in tissues, the oxidation- reduction (redox) ratio, defined as the ratio of FAD to NADH+FAD, has been used effectively as an indicator of metabolic activity as well for yielding more reliable information than measuring either NADH or FAD fluorescence intensity alone [3]. A redox ratio is a measure of aerobic glycolysis [16] to reflect the oxidized or reduced status of NADH and/or FAD. In tumors, these coenzymes are thought to exist in their reduced state, and thus a decrease in redox ratio is expected. A decrease in the redox ratio is thought to indicate an increase in the metabolic rate that it is typically observed in cancer cells possibly due to rapid cell division [4, 16]. Mitochondria are known to be one of the key regulators in cell death, which is now recognized as a complex continuum of mechanisms including the apoptotic pathways, necrosis, and autophagy [17]. Preclinical and clinical studies have shown that cell death can potentially be detected to provide an early indication of the success of therapy, providing prognostic information and guiding the course of subsequent treatments [18-20]. Specific, sensitive, rapid, reliable, and noninvasive detection of cell death in the clinic is urgently needed so that the early responses of tumors to drug treatment can be achieved. Currently, noninvasive in vivo imaging/detection techniques for the monitoring of cell death include magnetic resonance imaging (MRI), magnetic resonance spectroscopy (MRS), nuclear imaging [e.g., single-photon emission computed tomography and positron emission tomography (PET)], and optical imaging by combing with exogenous molecular markers such as the 40 kDa vesicle-associated protein, annexin V, and reporter probes that can be specifically cleaved by caspase 3 [18]. Ultrasound [21] and diffusion MRI [22] do not involve exogenous molecules, but indirectly measure cell death by measuring the scattering power of tissues and by measuring lipid droplets or apparent diffusion coefficient of water, respectively. The tissue scattering power change measured in ultrasound is due to nuclear condensation at the late stage of apoptotic process. In diffusion MRI, cell shrinkage due to apoptosis and/or cell swelling due to necrosis change tumor water diffusion [23]. Compared to ultrasound and diffusion MRI that detect cell death based on morphological characteristics of cells or tissues at the later stage of cell death, direct monitoring of the biochemical response of the mitochondrial dysfunction immediately after cell death is introduced using NADH and FAD intrinsic fluorescence, which may provide a new method for biomedical researchers to detect the early events of this process. Furthermore, it is clinically relevant to detect cell death in all its forms (e.g., apoptosis versus necrosis). The difference in the mitochondrial function between apoptosis and necrosis makes it possible to differentiate them using the NADH/FAD intrinsic fluorescence. This review aims to provide cell death detection information and update the study progress by NADH/FAD fluorescence spectroscopy and imaging. We describe recent advances in the following sections based on two divergent types of measurements, fluorescence intensity and time-resolved fluorescence (i.e., fluorescence lifetime). CELL DEATH MEASUREMENT BY NADH/FAD FLUORESCENCE INTENSITY Several studies have been conducted to assess the ability of optical techniques to monitor mitochondrial dysfunction during cell death in vitro [24-32] and in vivo [26, 27, 33, 34] by using different cell death inducers. Tables 1 and 2 summarize the results from these studies using different cell types (Table 1) or animal models used (Table 2), cell death stimuli, NADH and FAD intensity, and the redox ratio. NADH and FAD alone do not seem to have a trend to increase or decrease after treatment. Both increase and decrease

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in the fluorescence intensity were observed in response to different stimuli, the cell type, and the time point of measurement. In in vitro studies, Levitt et al. [30] observed an initial increase and then decrease in the fluorescence intensity of NADH and FAD prior to nuclear DNA fragmentation in cisplatin-induced apoptosis of human foreskin keratinocytes (HFKs) imaged by fluorescence microscopy. Similar trend was reported using flow cytometry and fluorospectroscopy in NADH fluorescence intensity of human lymphoid B cells and of yeast cells in response to campothecin-induced and H2O2/ONOO--induced apoptosis [24, 31], and in FAD fluorescence intensity of leukemic HL60 apoptotic cells following treatment with camptothecin, TNF- or -irradiation [28]. More specifically, Poot et al. [24] observed an increase in the intensity of NADH fluorescence at the so-called “non-apoptotic state” in which a slightly higher mitochondrial membrane potential (MMP) than controls was maintained. Besides, they observed a decrease in the NADH fluorescence at apoptotic state in which MMP was no longer maintained. The apoptotic cells that showed dynamic NADH fluorescence intensity (i.e., increase and then decrease) reported by Liang et al. [31] had a positive staining for Annexin V but not in propidium iodide staining. The increase and decrease in the FAD fluorescence reported by Wolbers et al. [28] was mainly seen in the early and late apoptotic subpopulation, respectively. Different from apoptosis, a very significant decrease in FAD fluorescence was seen in the necrotic subpopulation of the same study. Petit et al. [25] observed a monotonic decrease in the NADH fluorescence via CD95 antibody-induced apoptosis in Jurkat T cells before cells shrank and lost their capacity to exclude the vital stain of propidium iodide. They also found that the NADH fluorescence loss was correlated with the MMP dissipation after treatment. The similar decrease in the NADH fluorescence was observed by Lemar et al. [32] accompanied by a marked decrease of MMP, elevated intracellular ROS levels, and lower rate of oxygen consumption. Pogue et al. [26] observed a decrease in NADH fluorescence intensity immediately after benzoporphyrin derivative monoacid ring ‘a’ (BPD)-photodynamic therapy (PDT). On the other hand, Morbidelli et al. [29] observed an increase in the NADH fluorescence after simulated hypogravity-induced apoptosis of porcine aortic endothelial cells at single time point that most of hypogravity exposed cells already showed cell shrinkage and convolution of nuclear outline. An enhancement of the bound NADH according to the blue shift of the spectrum toward the shorter wavelength was accompanied by an increase in the fluorescence intensity of NADH. In terms of redox ratio in response to cell death in vitro, limited studies have been reported. However, there was a more consistent trend in that the redox ratio tended to increase afterwards. Levitt et al. [30] observed a decrease and then an increase in the redox ratio in the perinuclear ring formed by the redistribution of mitochondria and in the surrounding cytoplasmic areas of cells although the increased redox ratio stayed lower than the value before the redox ratio decreased in the perinuclear ring. Brewer et al. [27] observed that the increased redox ratio is correlated with apoptosis induced by N-(-hydroxyphenyl)retinamid (4-HPR) treatment of normal and simian virus-40 immortalized ovarian epithelial cells. Morbidelli et al. [29] observed increased or similar values in simulated hypogravity-exposed apoptotic cells for the ratio of FAD to free or bound NADH, respectively. The results of in vivo studies showed a similar pattern to the in vitro studies that NADH or FAD fluorescence intensity does not seem to have a trend to increase or decrease after cell death (apoptosis or necrosis) is introduced, but the redox ratio tends to increase consistently after treatments. Decreased NADH fluorescence intensity was observed in mouse leg muscle [26], female adult rhesus macaque ovary [27], and the skin of female Sprague-Dawley rats [34] after treated by BPD-PDT, oral contraceptive pill (OCP) and 4HPR/OCP combined treatment, and skin flap surgery, respectively. Conversely, increased NADH fluorescence intensity, although not

Reduced Nicotinamide Adenine Dinucleotide (NADH) Table 1.

Anti-Cancer Agents in Medicinal Chemistry, 2009, Vol. 9, No. 9

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Summary of the Change of Published NADH and FAD Fluorescence Intensity and Redox Ratio Defined as NADH/(NADH+FAD) Due to Various Cell Death Inducers In Vitro

Cell lines

Treatment

Cell death

NADH

FAD

Redox ratio

References

human lymphoid Bcell

Camptothecin

apoptosis

then

Jurkat T cell

CD95

apoptosis

RIF-1 cell

BPD-PDT

NOE, IOSE 261, IOSE 29

4-HPR

apoptosis

Human promyelocytic leukemic HL60

camptothecin, irradiation, TNF-/CHX, heat

apoptosis and necrosis

Porcine aortic endothelial cells

stimulated hypogravity

apoptosis

bound NADH

human foreskin keratinocytes (HFKs)

cisplatin

apoptosis

then

S. cerevisiae ATCC9080

H2O2, ONOO-


then (apoptosis); then significantly (necrosis)

Liang et al. 2007

Candida albicans (yeast cells)

DADS


Lemar et al. 2007

a

Poot Pierce 1999 a

Petit et al. 2001

Pogue et al. 2001

with apoptosis, with survival then (apoptosis), no change (necrosis)

then

Brewer et al. 2002

a

Wolbers et al. 2004

(FAD/free NADH), no change (FAD/bound NADH)

Morbidelli et al. 2005

then

Levitt et al. 2006

a: measurement was performed by flow cytometry

Table 2.

Summary of the Change of Published NADH and FAD Fluorescence Intensity and Redox Ratio Defined as NADH/(NADH+FAD) Due to Various Cell Death Inducers In Vivo

animal model

treatment

cell death

C3H/HeJ mice leg normal muscle

BPD-PDT

female adult rhesus macaques ovary

4-HPR, OCP, 4HPR+OCP

apoptosis

9L-glioma bearing rat

Pyro-2DG-PDT

necrosis

female rat skin

skin flap

necrosis

NADH

FAD

redox ratio

Pogue et al. 2001

(4-HPR), (OCP), (4-HPR+OCP)

(4-HPR), no change (OCP), (4HPR+OCP)

statistically significant, was observed in 4-HPR treatment of female adult rhesus macaque ovary [27]. Increase and decrease in the FAD fluorescence intensity were observed in 4-HPR treatment [27] and skin flap surgery [34], respectively. Finally, a consistent increase in the redox ratio was observed in female adult rhesus macaque ovary, 9L glioma-bearing rats, and the rabbit myocardium treated by 4HPR/OCP [27], pyropheophorbide-2-deoxyglucosamide (pyro2DG)-PDT [33], and heart infarction [34], respectively. However, it should be noted that the forms of induced cell death among these studies are different that necrosis was induced by pyro-2DG-PDT, but apoptosis was induced by 4-HPR/OCP and heart infarction. An increase in the redox ratio is not able to differentiate between apoptotic and necrotic forms of the cell death from these in vivo studies. CELL DEATH MEASUREMENTS BY NADH FLUORESCENCE LIFETIME Compared to fluorescence intensity measurement scheme, the fluorescence lifetime is a more sensitive probe of fluorophore microenvironment and offers a more sensitive discrimination of free and enzyme bound forms of NADH and FAD. For example, the binding-induced shift of the NADH fluorescence emission spectrum

reference

(4-HPR, OCP, 4HPR+OCP)

Brewer et al. 2002 Zhang et al. 2004 Mokry et al. 2007

is ~10 to 20 nm that is small compared with the width of the NADH emission spectrum (~150 nm) [35]. In contrast, up to 10 times of the fluorescence lifetime change was observed in response to NADH binding [36-38]. Intracellular NADH exists in dynamic pools of both free NADH and those bind to various dehydrogenases. The fluorescence decay is thus a multi-exponential function attributing to both free and multiple bound components [38, 39]. Several studies have shown that the free NADH pool has a short lifetime of ~0.4 ns and bound NADH pool has a wider range of longer lifetime, ~1 to 8 ns [37-39]. Different from NADH, the bound FAD has a short lifetime component and the free FAD has a long lifetime component [40]. For the simplicity of multiple lifetime parameters involved and some commercial analytical software limitation, two-component exponential function F(t) is commonly used to fit the measured NADH fluorescence decay curve so that F (t ) = a1e t 1 + a2 e t 2 , where 1 and 2 represent the short and long lifetime, respectively, and a1 and a2 are the corresponding relative amplitudes or contributions [35, 41]. a1 to a2 ratio has been often used to represent the ratio of the contribution between free and bound NADH. The mean fluorescence lifetime m is defined as


m=(a11 + a22 ). Same as the NADH fluorescence decay curve, FAD fluorescence decay curve can be also fit to a two-component exponential function to obtain lifetime parameters similar to those of NADH. In the following paragraphs of this review, we refer the lifetime parameters 1, 2, a1, a2, and m only specific to the NADH fluorescence. There are only few studies available on the dynamic redistribution of NADH fluorescence lifetime during cell death. Pogue et al. [26] observed no lifetime change in vivo in the mouse leg muscle after BPD-PDT treatment, which likely induced necrosis. Wang et al. [42] observed initially significant increase (up to 2 ns) and then decrease in the mean NADH fluorescence lifetime (m) in staurosporine (STS)-induced apoptosis, but no lifetime change was observed in H2O2-induced necrosis in HeLa and 143B cells. This increased NADH fluorescence lifetime is suggested to reflect early mitochondrial responses of STS-induced apoptosis because it appeared before the phosphatidylserine exposure (annexin V stain positive < 4%) and caspase-3 activation. It remains unclear whether these observations are related to NADH binding to different proteins in mitochondria for more efficient generation of ATP required for the support of apoptotic process. Further studies are on going in our laboratories to reveal the relationship between NADH fluorescence lifetime dynamics, mitochondrial membrane potential, mitochondrial respiration state, and ATP production and/or the maintenance of mitochondrial function. To successfully apply NADH/FAD fluorescence lifetime in future detection of cell death, it is essential to understand the origins of fluorescence lifetime changes due to binding mechanism during physiological and/or metabolic perturbation. Wakita et al. [38] reported that the mean lifetime of NADH fluorescence stayed constant regardless of numerous perturbations including induction of active ATP production (State 3), resting (State 4) respiration, and uncoupled conditions, and conditions with various lactate/pyruvate ratios. They observed that the NADH fluorescence intensity, as reported in many previous studies, responded to these perturbations so that fluorescence intensity decreased at State 3 respiration, increased at State 4 respiration, and increased with increased lactate/pyruvate ratio, and further increased in anaerobic state as glu-

Wang et al.

tamate was used as a substrate. Other studies to report NADH lifetime changes due to various inhibitors of the cellular metabolism are limited to Complex IV inhibitor potassium cyanide (KCN) [41] and sodium cyanide (NaCN) [35], uncoupler carbonyl cyanide-3chloro-phenylhydrazene (CCCP) [35], and glycolysis inhibitor 3bromopyruvate [43]. Bird et al. [41] observed ~20 to 40% increased a1/a2 within 10 minutes after KCN administration in MCF10A human breast cells. Wu et al. [35] observed up to 16% increased a1/a2 due to NaCN treatment and up to 13% decreased a1/a2 due to CCCP uncoupler in cancerous SiHa cells although the treatment time was not reported. Skala et al. [43] reported a less than 4% increase in 2 and decrease in a2 depending on the cell confluence after 3bromopyruvate treatment of human breast cancer cells MCF10A. We studied NADH fluorescence lifetime dynamics (up to 24 hours after treatment) in response to various respiratory inhibitors including Complex I inhibitor rotenone, Complex III inhibitor antimycin A, Complex IV inhibitor KCN, Complex V inhibitor oligomycin, and glycolysis inhibitor 2-deoxy D-glucose (2-DG) in normal skin fibroblasts. No significant change of lifetime parameters (a1, a2, a1/a2,, and m) except that perturbed by KCN. Fig. (1) illustrates two-photon fluorescence lifetime microscopic images of normal skin fibroblasts before and after treatment with KCN [Fig. (1a)] and oligomycin [Fig. (1b)] at the first 10 minutes and at 55 to 65 minutes. Mean lifetime m is color coded from 0.5 ns (red color) to 2 ns (blue color). The cells treated with KCN at the first 10 minutes turned to more yellowish color than control and the lifetime returned back to similar to controls at 55 to 65 minutes. No change in lifetime encoded color was seen in olygomycin treated cells. Fig. (2) plots the average results (n=3 to 5) of m [Fig. (2a)) and a1/a2 [Fig. (2b)] as a function of time before and up to 24 hours after treatment with KCN, oligomycin, and 2-DG, respectively. The decrease in m and increase in a1/a2 at the first 10 minutes after KCN treatment agree with previous published studies using cyanide as inhibitors by Wu et al. [35] and Bird et al. [41]. However, both m and a1/a2 returned back to the values of controls after ~1 hour. Apparently, the difference of the change between the NADH fluorescence lifetime and intensity affected by metabolic perturbation indicates that these changes of both signals are coming from different origins. Studies by Blinova et al. [13, 14] suggested that the

Fig. (1). The mean lifetime (m) of NADH fluorescence in normal fibroblasts before and after treatment with (a) KCN and (b) olygomycin at the first 10 minutes and 55 to 65 minutes. The lifetime is color-coded from 0.5 ns (red color) to 2 ns (blue color). NADH fluorescence is two-photon excited at 740 nm and collected at 450±40 nm.

Reduced Nicotinamide Adenine Dinucleotide (NADH)

NADH associated with Complex I significantly contributes to the overall mitochondrial NADH fluorescence intensity so that it provides an explanation for the close correlation between mitochondrial NADH fluorescence intensity and the metabolic state observed in many previously published perturbation studies. On the other hand, the NADH fluorescence lifetime is not only attributed to NADH binding to Complex I, but also its binding to multiple dehydrogenases in addition to free NADH in mitochondria and/or cytoplasm.

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scheme revealed both increased and decreased NADH/FAD fluorescence intensity, but more consistently increased redox ratio was seen after the induction of cell death. Significant NADH/FAD fluorescence loss tends to correlate with the MMP dissipation, late stage of apoptosis, and necrosis. Limited studies based on lifetime measurement scheme observed significant lifetime increase of NADH fluorescence in the beginning of STS-induced apoptosis, but no lifetime change was observed in necrosis induced by H2O2- and PDT, respectively. The fact that the measurement of NADH fluorescence lifetime responded differently from that of NADH fluorescence intensity to metabolic perturbations is likely due to the contribution of the NADH bindings. The NADH fluorescence intensity is mainly attributed to Complex I binding but NADH fluorescence lifetime is attributed to multiple protein bindings. The understanding of the actual binding site and mechanism for a specific death pathway is important for the successful application of NADH/FAD fluorescence lifetime in the future detection of cell death in vivo. ACKNOWLEGMENTS We acknowledge the support of Imaging Core Facility of National Yang-Ming University. This work is supported by the “Aim for Top University Plan” from the Ministry of Education of Taiwan, and grants NSC 94-2321-B-010-004-YC and NSC 95-2112-M-010002 from the National Science Council of Taiwan. REFERENCES [1] [2]

[3] Fig. (2). (a) The mean lifetime (m) and (b) a1 /a2 ratio change of the NADH fluorescence as a function of time before (time = 0) and after treatment with various inhibitors including KCN, olygomycin, and 2-DG.

Finally, studies in precancerous epithelial cells in vivo [43, 44] and human mesenchymal stem cells (hMSCs) in vitro [45] have shown that NADH fluorescence lifetime is associated with the change of the transition from anaerobic glycolysis to aerobic metabolism. In a hamster cheek pouch model of oral cancer in vivo, Skala et al. [43, 44] observed decreased 2 and a2 of the NADH fluorescence lifetime, thus corresponding to decreased m and increased a1/a2, in low and high-grade precancerous epithelial cells where cells tend to metabolize by glycolysis rather than oxidative phosphorylation. Guo et al. [45] observed an increased m and a decreased a1/a2 during stem cell differentiation when cellular metabolism gradually changed from glycolysis to aerobic metabolism. The NADH lifetime was observed to be sensitive to hypoxia environment as well. A number of studies of cell culture [41, 43, 46] and tissue slices [39] have shown a decrease in 1, 2 and a2, which corresponds to a decrease in m and an increased a1/a2 in hypoxia. These results indicate that decreased m and increased a1/a2 are correlated with anaerobic glycolysis of precancerous tissues and stem cells, and hypoxic environment. Although the actual change of NADH binding sites during this transition is unknown, it warrants further studies to understand the sources underlying these changes that may help explain the origins of previously detected increase of the NADH fluorescence lifetime in STS-induced apoptosis. SUMMARY NADH/FAD fluorescence lifetime has the potential to noninvasively detect cell death in vivo at its early stage and to differentiate different forms of death (e.g., apoptosis versus necrosis) due to the various biochemical reactions of the mitochondrial dysfunction at different death pathways. However, these biochemical reactions can be complicated. Up to date, studies based on intensity measurement

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Received: June 17, 2009

Revised: June 02, 2009

Accepted: July 12, 2009

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