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Published as doi: 10.1096/fj.08-112375.
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(The FASEB Journal. 2008;22:3878-3887.)
© 2008 FASEB

Vitamin A depletion causes oxidative stress, mitochondrial dysfunction, and PARP-1-dependent energy deprivation

Haw-Jyh Chiu*, Donald A. Fischman{dagger} and Ulrich Hammerling*,1

* Memorial Sloan-Kettering Cancer Center, Immunology Program, New York, New York, USA; and

{dagger} Department of Cell and Developmental Biology, Weill Medical College of Cornell University, New York, New York, USA

1 Correspondence: Ulrich Hammerling, Memorial Sloan-Kettering Cancer Center, Immunology Program, 1275 York Ave., New York, NY 10065, USA. E-mail: u-hammerling{at}ski.mskcc.org


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
A significant unresolved question is how vitamin A deprivation causes, and why retinoic acid fails to reverse, immunodeficiency. When depleted of vitamin A, T cells undergo programmed cell death (PCD), which is enhanced by the natural competitor of retinol, anhydroretinol. PCD does not happen by apoptosis, despite the occurrence of shared early events, including mitochondrial membrane depolarization, permeability transition pore opening, and cytochrome c release. It also lacks caspase-3 activation, chromatin condensation, and endonuclease-mediated DNA degradation, hallmarks of apoptosis. PCD following vitamin A deprivation exhibits increased production of reactive oxygen species (ROS), drastic reductions in ATP and NAD+ levels, and activation of poly-(ADP-ribose) polymerase (PARP) -1. These latter steps are causative because neutralizing ROS, imposing hypoxic conditions, or inhibiting PARP-1 by genetic or pharmacologic approaches prevents energy depletion and PCD. The data highlight a novel regulatory role of vitamin A in mitochondrial energy homeostasis.—Chiu, H.-J., Fischman, D. A., Hammerling, U. Vitamin A depletion causes oxidative stress, mitochondrial dysfunction, and PARP-1-dependent energy deprivation.


Key Words: lymphocytes • mitochondrion • reactive oxygen species • bioenergetics


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
VITAMIN A (RETINOL) IS ESSENTIAL for a diversity of physiological processes, including vision, embryonic development, skin differentiation, spermatogenesis, and immune system function (1) . What separates the last 2 from the rest is that these are dependent on native retinol (ROL); retinoic acid (RA) can only incompletely substitute for ROL, if at all, suggesting the presence of a mechanism distinct from the classic gene transactivation paradigm attributed to RA (2 , 3) . Virtually all tissues take up vitamin A, protect it from degradation by cellular ROL binding protein, convert it to retinyl-ester for storage, and, on demand, recover free ROL by ester hydrolysis. Moreover, ROL is biosynthetically converted to a number of metabolites other than RA, and these do not play a direct role in the nucleus. Among them are 14-hydroxy-retro-retinol and 13,14-dihydroxy-retinol, both prevalent in many tissues and cells, including some not known to synthesize RA (2 3 4) . Moreover, ROL-dependent functions appear to be evolutionarily older than transcriptional regulation by RA, which first evolved at the invertebrate/vertebrate boundary. Invertebrates have the capacity to metabolize ROL, as shown by biosynthesis of the above-mentioned hydroxylated retinoids, as well as anhydroretinol (AR), in Spodoptera frugiperda (2) .

These considerations raise the question of what biological functions these ROL metabolites, and indeed ROL itself, do perform. A partial answer was obtained by the discovery that they serve as essential cofactors in signaling pathways controlling proliferation and survival of B lymphoblasts, activated T lymphocytes, murine fibroblasts, and many other cell types (2 3 4 5 6 7 8 9) . AR is a natural, cellular ROL derivative that antagonizes the survival-promoting properties of ROL (6 , 8 9 10) . The biochemical basis for AR function is still unknown, but it may be linked to its capacity to displace ROL from common receptor sites we identified on serine/threonine kinases (11 12 13 14 15) . These receptor sites have been localized to regulatory domains of the Raf and protein kinase C (PKC) families, specifically the zinc-finger domains that also harbor their activation centers. Although irrelevant for classic activation pathways via the diacylglycerol second messenger, these sites must be occupied by ROL for efficient activation by nonclassic stress signals, notably reactive oxygen species (ROS) (12 , 14 , 15) . When left empty or occupied by AR, we surmise that ROL-dependent survival signals are disrupted, leading to rapid cell death.

Although confirming the previous notion that the mode of cell death resulting from vitamin A deprivation is characterized by increased ROS formation (16) , the present studies reveal mitochondrial membrane depolarization and rapid loss of plasma membrane integrity as additional consequences, but no overt activation of caspases or chromatin condensation, both hallmarks of apoptotic programmed cell death (17) . Significant decreases in cellular levels of ATP and NAD+ point to the mitochondrion as the principal site of injury, but vitamin A deprivation also causes the activation of poly-(ADP-ribose) polymerase (PARP) -1. PARP-1, the best-characterized member among the PARP family of nuclear proteins, catalyzes the NAD+-dependent synthesis and attachment of ADP-ribose units to {gamma}-carboxyl groups of Glu residues of acceptor proteins (18 , 19) . These include PARP-1 itself, histones, topoisomerases I and II, DNA helicases, single-strand base repair and base-excision repair factors, and various transcription factors (18 , 19) . In addition to its role as a sensor of DNA damage (20) , PARP-1 is also involved in regulation of transcription, cell cycle, and cell death (21) . Overactivation of PARP-1 under cellular stress, with the resultant formation of high levels of PAR and PAR-modified proteins, has recently been shown to cause caspase-independent cell death in models of ischemia-reperfusion injury, diabetes, inflammatory-mediated injury, and neurotoxicity (22 23 24 25) . Studies have suggested that mechanisms of PARP-1-induced cell death include energy failure, due to rapid utilization of NAD+ and ATP, transcriptional derangement, and PAR-mediated signaling to the mitochondria (24 , 26 27 28 29) . Using pharmacologic and genetic approaches, we now demonstrate that cellular deprivation of vitamin A, in addition to antagonizing normal ROL-dependent survival signals by AR, produces unremitting oxidative stress, mitochondrial dysfunction, PARP-1 activation, and eventually necrotic cell death from bioenergetic crisis. It is noteworthy that these pathological responses spotlight the mitochondrion as a hitherto unsuspected target of vitamin A regulation.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Reagents
ROL, (±)-6-hydroxy-2,5,7,8-tetramethylchromane-2-carboxylic acid (Trolox), carbonyl cyanide 3-chlorophenylhydrazone (CCCP), myxothiazol, retinoic acid p-hydroxyanilide (4-HPR, fenretinide), 1,5-isoquinolinediol (DHIQ), and formaldehyde were purchased from Sigma-Aldrich (St. Louis, MO, USA). Propidium iodide, tetramethylrhodamine ethyl ester perchlorate (TMRE), and 5-(and-6)-carboxy-2',7'-DCF diacetate (carboxy-DCFDA) were obtained from Invitrogen (Carlsbad, CA, USA). Mouse anti-human PAR was purchased from BD Biosciences (San Jose, CA, USA). Mouse anti-human Fas (CH11) was obtained from Upstate (Temecula, CA, USA). CD90 (Thy1.2) MicroBeads were purchased from Miltenyi Biotec (Auburn, CA, USA). Boc-Asp(OMe)-CH2F (Boc-D-FMK) was obtained from EMD Biosciences (San Diego, CA, USA).

Animals and cells
C57BL/6 mice were obtained from Jackson Labs (Bar Harbor, ME, USA). Male mice were used for all experiments. The animals were maintained and treated according to guidelines of the Animal Health Services of Memorial Sloan-Kettering Cancer Center (MSKCC). All mouse experiments were performed under a protocol approved by the Institutional Animal Care and Utilization Committee of MSKCC. Jurkat T cells were purchased from American Type Culture Collection (Manassas, VA, USA) and grown in RPMI 1640 (Invitrogen) supplemented with 10% heat-inactivated FBS (Omega Scientific, Tarzana, CA, USA) and maintained at 37°C with 5% CO2. For hypoxia experiments, cells were exposed to hypoxia (37°C, 1% O2, 5% CO2, 94% N2) with medium equilibrated overnight with hypoxic gas mixture in a modular incubation chamber (Billups Rothenberg, Del Mar, CA, USA).

Cell viability
Jurkat T cells, CD90 (Thy1.2)+ T cells purified by positive immunomagnetic selection (Miltenyi Biotec) from spleens of C57BL/6 mice, or wild type and PARP-1–/– Mouse embryonic fibroblasts (MEFs) were cultured in serum-free TLB medium (RPMI 1640, 20 µg/ml transferrin, 5 µM linoleic acid, 0.12% bovine serum albumin) containing AR (with or without various inhibitors) for 1–24 h in 96-well plates. Cell Proliferation Reagent WST-1 (Roche, Mannheim, Germany) was added to Jurkat T, Thy1+, or MEF cells for an additional 2 h. The absorbance (Abs450 nm to Abs690 nm) was determined using a 96-well reader (Molecular Dynamics, Sunnyvale, CA, USA). Viability was expressed as a percentage of control untreated cells. Separately, Jurkat T cells were collected, washed with ice-cold PBS, and labeled with 10 µg/ml propidium iodide for 5 min at room temperature. Propidium-iodide-positive cells were acquired on a FACScan (Becton Dickinson, San Jose, CA, USA) and analyzed using FlowJo (TreeStar, San Carlos, CA, USA).

Apoptosis assays
Apoptosis was assayed via flow cytometric detection of cleaved caspase-3 and TUNEL staining. Jurkat T cells incubated in 4 µM AR for 6 h were washed in ice-cold PBS, fixed in 3% formaldehyde for 10 min at 37°C, and permeabilized in 90% ice-cold methanol for 30 min on ice. Cells were incubated for 60 min at 25°C with 1:500 cleaved-caspase-3 polyclonal antibody, followed by Alexa-Fluor488-labeled anti-rabbit immunoglobulin G (IgG; 1:1000) for 30 min at 25°C. For TUNEL assay, the in situ Cell Death Detection Kit (Roche) was used. Briefly, cells were washed in ice-cold PBS, fixed in 2% formaldehyde for 60 min at 25°C, permeabilized in 0.1% Triton X-100 in 0.1% sodium citrate, and incubated in TUNEL reaction mixture for 60 min at 37°C in the dark. Caspase-3 antibody binding and labeling on DNA strand breaks were analyzed on a FACScan (Becton Dickinson) and analyzed using FlowJo (TreeStar). At least 10,000 cells of each sample were analyzed.

Mitochondrial membrane potential
Mitochondrial membrane potential was measured by flow cytometry using TMRE, a cationic membrane permeable reagent (30) . Cells were loaded with TMRE for the last 30 min of cell treatment, and then washed in ice-cold PBS; cellular staining was measured on a FACScan (Becton Dickinson), and data were analyzed using FlowJo (TreeStar). At least 10,000 cells of each sample were analyzed.

ATP and NAD+ measurement
After treatment, Jurkat T cells were washed with cold PBS and collected in separate tubes. For ATP measurement, samples were lysed in boiling buffer, 100 mM Tris with 4 mM EDTA (pH 7.75), for 5 min and centrifuged for 3 min at 1000 g. ATP in the supernatant was quantitatively measured by the ATP Bioluminescence Assay Kit CLS II (Roche). For NAD+ measurement, samples were lysed by homogenization and centrifuged at 14,000 rpm for 5 min. NAD+ in the supernatant was quantified by the NAD+/NADH Quantification Kit (Medical Biological Laboratories, Woburn, MA, USA).

Cellular ROS measurement
ROS production was measured by flow cytometry using 5-(and-6)-carboxy-2',7'-DCF diacetate, a membrane permeable and oxidant-sensitive reagent. Cells were treated with this reagent 30 min before the end of a time point, washed in ice-cold PBS, and analyzed on a FACScan (Becton Dickinson) and analyzed using FlowJo (TreeStar). At least 10,000 cells of each sample were analyzed.

PAR detection
Jurkat T cells incubated in 4 µM AR for 4 h, were washed in ice-cold PBS, fixed in 3% formaldehyde for 10 min at 37°C, and permeabilized in 90% ice-cold methanol for 30 min on ice. Cells were incubated for 60 min at 25°C with 1:500 PAR polyclonal antibody, followed by Alexa-Fluor488-labeled anti-rabbit IgG (1:1000) for 30 min at 25°C. PAR antibody binding was analyzed on a FACScan (Becton Dickinson) and analyzed using FlowJo (TreeStar). At least 10,000 cells of each sample were analyzed.

Statistical analyses
All values are expressed as means ± SE, unless indicated otherwise. Data were analyzed using Student’s t test. Results were considered statistically significant at P < 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Vitamin A depletion, as well as antagonism elicited by anhydroretinol, causes caspase-independent cell death
T-cell blasts generated by stimulation with anti-CD3 antibody and cultured in vitro are dependent on ROL for survival in serum-free culture medium. When the ROL levels drop below the physiological range of 1–2 µM, blasts perish within 24 h (Fig. 1A ). AR accelerates this process. In keeping with its function as a mutual reversible inhibitor of vitamin A, AR-induced cell death is reversed by ROL in a dose-dependent manner (Fig. 1A ). Treatment with AR for 24 h, under serum-free conditions, causes dose-dependent death of Jurkat cells (Fig. 1B, C ), similar primary T-cell blasts. The cytotoxic effect of AR is attenuated when progressively increasing concentrations of ROL are added simultaneously to a maximum of 2 µM (Fig. 1B, C ). To determine the degree of latent damage and the time point when ROL is still capable of rescue, Jurkat cells were cultured in AR for different time periods before being returned to ROL-containing medium. Cells withstand AR for limited time periods, depending on dose, but reach a point, measured in hours, when they become irreversibly committed to cell death (Fig. 1D ).


Figure 1
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  		Figure 1. ROL deprivation causes a time- and dose-dependent decrease in viability of primary Thy1.2+ mouse lymphoblasts and Jurkat T cells. A) Primary Thy1.2+ mouse lymphocytes were activated by plate-bound anti-CD-3 antibody and after 24 h incubated in serum-free medium alone, in the presence of 1–3 µM AR alone, or in combination with 0.03–2 µM ROL for an additional 24 h. Cell viability was assayed by spectrophotometrical quantification of formazan formation. A representative of 3 independent experiments is shown. B) Jurkat T cells were incubated in serum-free medium with 0.5–4 µM AR alone or in combination with 2 µM ROL for 24 h and assayed for cell viability by spectrophotometrical quantification of formazan formation. Note that AR-induced cell dysfunction was associated with decreased formazan formation, and ROL significantly reversed the cytotoxic effect of AR. Data represent means ± SE of 3–4 independent experiments. C) Jurkat T cells incubated with 4 µM AR for 24 h were assayed for cell viability by flow cytometrical analysis of light side-scattering or propidium iodide uptake. Note that AR treatment caused increased light side-scattering and propidium iodide uptake. A representative of 3 independent experiments is shown. D) Jurkat T cells incubated with medium or 2 µM AR for 1 to 3 h were washed and cultured for an additional 24 h in assay medium with or without 2 µM ROL, and cell viability assayed by flow cytometric analysis of propidium iodide uptake. Note cell survival was dependent on the amount of time cells were exposed to AR. Incubation with ROL after treatment with AR led to significant increase in cell survival. Data represent means ± SE of 3 independent experiments. *P < 0.01 vs. untreated control. **P < 0.01 vs. AR treatment alone.

To identify possible signaling pathways affected by vitamin A deprivation, we analyzed the mechanism of cell death. In contrast to Fas-mediated apoptosis, no significant caspase-3 activation or DNA fragmentation is detected following 6 h of AR treatment (Fig. 2A, B ). Concordantly, AR-induced cell death is not inhibited by the irreversible pan-caspase inhibitor Boc-D-FMK (Boc-Asp(OMe)-CH2F) (Fig. 2C ). These results suggest that cell death proceeds through a nonapoptotic pathway, consistent with previous observations by Chen et al. (16) of a caspase-independent cell death in B lymphoblastoid cells. Similar to AR-induced PCD, cell death of T cells caused by ROL deprivation displays minimal signs of DNA damage (9) and is caspase-3 independent (data not shown).


Figure 2
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  		Figure 2. AR mediates a nonapoptotic, caspase-independent cell death. A, B) Apoptosis was measured by flow cytometric detection of cleaved-caspase-3 (A) and TUNEL (B) staining in Jurkat T cells incubated with 4 µM AR or 10 ng/ml Fas antibody (CH11) for 6 h. Note that AR-treatment did not result in increased cleaved-caspase 3 or TUNEL staining. A representative experiment of 3 independent experiments is shown. C) Cell viability was assayed in Jurkat T cells incubated with medium control, 0.5–4 µM AR, or 1 ng/ml Fas antibody (CH11), in the absence (control) or presence (+caspase inhibitor) of the broad-base caspase inhibitor, Boc-D-FMK. Note that inhibition of caspases had no effect on AR-induced cytotoxicity. Data represent means ± SE of 3–4 independent experiments. *P < 0.01 vs. untreated control. **P < 0.01 vs. DMSO pretreatment.

Mitochondrial membrane potential and cellular ATP levels are decreased following treatment with AR
Mitochondrial permeability transition is widely accepted as an important regulator of cell death (31) . Disruption of mitochondrial membrane potential ({Delta}{Psi}m) can lead to increased production of ROS, leakage of proapoptotic proteins into the cytoplasm and nucleus, and decreased ATP production. To determine whether mitochondrial parameters are altered during AR-induced cell death resulting from ROL depletion, we measured ROS by the vital stain, dichloro-fluorescein-diacetate (DCFDA), {Delta}{Psi}m by the cationic potential-sensitive dye, tetramethylrhodamine-ethyl-ester (TMRE), which accumulates in intact mitochondria (30) , intracellular ATP by the luciferase assay, and NAD+ levels by a colorimetric enzyme cycling reaction. Within 4 h in ROL-deprived culture cell injury is evident: there is a drop in {Delta}{Psi}m, indicated by the modest increase of Jurkat cells with low TMRE fluorescence compared to ROL-sufficient cells, but this is dramatically increased in cells cultured with AR (Fig. 3A ). As expected, the decrease in {Delta}{Psi}m in both ROL-depleted and AR-treated cells, relative to ROL-sufficient cultures, is paralleled by rapid declines in the steady-state levels of cellular ATP and NAD+ (Fig. 3B ). Our results suggest that when Jurkat cells run out of ROL or are acutely deprived by blockade of endogenous ROL by AR, their cellular ATP levels drop steeply. ATP deprivation interferes with the execution phase of apoptosis requiring ATP (32 33 34) . Instead, cell death occurs by necrosis.


Figure 3
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  		Figure 3. AR causes mitochondrial membrane depolarization and decreased cellular levels of ATP and NAD+. A) Mitochondrial membrane potential was measured by flow cytometric analysis of TMRE fluorescence in Jurkat T cells after 4 h incubation in medium (control), 2 µM ROL, or 4 µM AR. A representative experiment of 6 independent experiments is shown. B) Jurkat T cells incubated with medium (untreated), 2 µM ROL, or 2 µM AR for 2–6 h were collected and analyzed for cellular ATP and NAD+ levels. Initial [ATP] = 129.60 ± 10.09 pM/µg protein; initial [NAD+] = 1814.04 ± 357.66 ng/mg protein. Note that AR treatment caused a rapid decrease in cellular levels of ATP and NAD+. Data represent means ± SE of 3–4 independent experiments. *P < 0.01 vs. untreated control.

Anhydroretinol activates PARP-1, but pharmacological blockade or gene disruption of PARP-1 protects from cell death
PARP-1 overactivation during cellular injury has been shown to cause ATP and NAD+ depletion and mitochondrial dysfunction, leading to caspase-independent cell death (24 , 26 27 28 29) . Although gradual ROL deprivation by removing the exogenous source is insufficient to cause PARP-1 activation in Jurkat cells, acute deprivation by blocking endogenous ROL with AR causes a rapid accumulation of PAR. Simultaneous treatment with ROL and AR completely prevents this response (Fig. 4A ). Pharmacological inhibition of PARP-1 by the potent and specific inhibitor DHIQ (35) suppresses PAR adduct formation (Fig. 4A ) and significantly improves cell survival following otherwise lethal doses of AR, most cells remaining viable 24 h posttreatment (Fig. 4B ). Similarly, MEFs with germ-line disruption of the PARP-1 gene are largely protected against AR-induced cytotoxicity compared to wild-type cells (Fig. 4C ). Furthermore, DHIQ prevents the rapid and significant loss of cellular levels of ATP and NAD+ following AR treatment (Fig. 4D, E ). These data suggest that PARP-1 overactivation and the subsequent PARP-1-dependent cellular ATP and NAD+ depletion exacerbate the detrimental effects of ROL deprivation.


Figure 4
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  		Figure 4. PARP-1 activation by AR and protection against cell death in PARP-1–/– MEFs and by PARP-1 inhibitor, DHIQ, by preventing depletion of cellular ATP and NAD+. A) Flow-cytometric measurement of cell death of Jurkat T cells incubated with medium (control), 2 µM ROL, or 4 µM AR alone or in combination with 2 µM ROL or 150 µM DHIQ for 4 h were collected and analyzed for PAR levels by flow cytometry. A representative experiment of 3 independent experiments is shown. B) Cell viability was analyzed 24 h after incubation of Jurkat T cells in medium (control) or 0.5–2 µM AR alone or in the presence of DHIQ. Data represent means ± SE of 4–5 independent experiments. *P < 0.01 vs. untreated control. **P < 0.01 vs. DMSO pretreatment. C) PARP-1+/+ or PARP-1–/– MEFs incubated with medium (control) or 1–2 µM AR for 24 h were analyzed for cell viability by spectrophotometrical quantification of formazan formation. Data represent means ± SE of 6 independent experiments. *P < 0.03 vs. untreated control. **P < 0.03 vs. AR treatment alone. D, E) Jurkat T cells incubated for 2–6 h with 2 µM AR alone or together with 150 µM DHIQ were assayed for intracellular levels of (D) ATP and (E) NAD+. Data represent means ± SE of 3–4 independent experiments. *P < 0.01 vs. untreated control. **P < 0.01 vs. DMSO pretreatment.

Hypoxia exposure abrogates ROS production, mitochondrial membrane depolarization, and cell death following AR treatment of Jurkat cells
ROS and dissipation of the mitochondrial membrane potential are common initiators of cell death (31) . Early consequences of exogenous ROL deprivation or AR antagonism are the production of ROS (Fig. 5A ) (16) . To identify the source of ROS, Jurkat cells were first incubated with either CCCP, an uncoupler of oxidative phosphorylation that abolishes the mitochondrial membrane proton gradient, or myxothiazol, an inhibitor of the Rieske iron-sulfur protein in complex III of the electron transport chain (36) . Both compounds prevent ROS formation following ROL depletion or AR treatment (Fig. 5B ), similar to reports showing significant decrease in hyperglycemia-induced ROS formation by these agents in bovine aortic endothelial cells (37 , 38) , pointing to the mitochondria as the source of ROS during ROL deprivation. Consistent with its role as a potent ROS scavenger, the vitamin E derivative Trolox significantly reduces AR-induced cellular ROS formation, reverses the loss of ATP and NAD+ (Fig. 5C ), and in large part prevents cell death (Fig. 5D ). It is significant that ROS production, like cell death, is prevented when ROL is present simultaneously with AR (Fig. 5A ). Our data suggest that when mitochondrial dysfunction is prevented and ROS levels are normalized, cell death due to AR exposure is largely avoided.


Figure 5
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  		Figure 5. Trolox prevents increases in cellular ROS and cell death following AR treatment. A) Jurkat T cells incubated for 2 h with medium, 2 µM ROL, 100 µM Trolox, or 4 µM AR alone or together with ROL or Trolox, were collected and assayed for cellular ROS production by flow cytometric detection of oxidized-CH2DCFDA. A representative experiment of 4 independent experiments is shown. B) Jurkat cells incubated for 2 h with 4 µM AR alone or together with 0.5 µM myxothiazol, 10 µM CCCP, or 100 µM Trolox were collected, and ROS production was assayed by flow cytometric detection of oxidized-CH2DCFDA. Note that, like Trolox, CCCP and myxothiazol significantly decreased AR-induced increase in ROS production. A representative experiment of 3 independent experiments is shown. C) Jurkat T cells incubated for 2–6 h with 2 µM AR alone or together with 100 µM Trolox were assayed for intracellular levels of ATP and NAD+. Data represent means ± SE of 3–4 independent experiments. D) Jurkat T cells incubated for 24 h with medium (control) or 1–4 µM AR alone or together with 1–100 µM Trolox were assayed for viability. Note that Trolox protected Jurkat cells against AR-induced cell death in a dose-dependent manner. Data represent means ± SE of 5 independent experiments. *P < 0.01 vs. untreated control. **P < 0.01 vs. ethanol pretreatment.

As migratory cells, lymphocytes evolved strategies to cope with different oxygen tensions, including the acute hypoxia they endure while entering ischemic, inflamed tissues during infection (39 40 41) . To investigate the effects of ROL antagonism by AR during hypoxia, Jurkat cells were cultured in 1% oxygen. We observe that hypoxic conditions significantly ameliorate AR-induced cellular injury (Fig. 6A ). Furthermore, hypoxia decreases cell death caused by the synthetic retinoid, fenretinide, but not by Fas antibody (CH11) (Fig. 6B, C ), which is known to cause classical caspase-dependent apoptosis in Jurkat cells (42 43 44) . Fenretinide has previously been shown to cause cell injury by increasing ROS production by mitochondrial-dependent pathways (42 , 43 , 45 , 46) . Interestingly, hypoxia, unlike DHIQ, completely inhibits ROS production in Jurkat cells following withdrawal of exogenous ROL or blockade of endogenous ROL by AR (Fig. 7A, B ). Conversely, DHIQ and AR synergize in ROS generation. These data suggest that protection from overproduction of ROS by hypoxia, like Trolox, also protects against cell death, pointing to ROS as the likely primary upstream initiator of cell injury. Whereas poly(ADP-ribosyl)ation contributes to loss of cell viability, the action of PARP occurs downstream of mitochondrially derived ROS and represents a key process in cell death execution. When PARP is blocked by DHIQ, or dysfunctional due to gene disruption, cell viability is not in jeopardy.


Figure 6
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  		Figure 6. Hypoxia protects against cell injury by AR and fenretinide but not Fas antibody. Jurkat T cells incubated for 24 h with medium or 0.5–5 µM AR, 0.125–1 µM fenretinide, or 0.01–1 ng/ml Fas antibody under normoxic or hypoxic conditions were assayed for cell viability by spectrophotometrical quantification of formazan formation in viable cells. Note that hypoxia protected Jurkat cells against cell death by AR and fenretinide but not Fas antibody. Data represent means ± SE of 3 independent experiments. *P < 0.01 vs. untreated control. **P < 0.05 vs. treatment under normoxic conditions.


Figure 7
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  		Figure 7. Hypoxic exposure prevents accumulation of ROS in Jurkat T cells following AR treatment. A) Jurkat T cells were incubated for 1–6 h in ROL-free medium, or medium supplemented with 2 µM ROL, 2 µM AR, or 100 µM Trolox, alone or in the combinations indicated, under normoxic or hypoxic conditions. Cells were assayed for ROS formation by flow cytometric detection of oxidized carboxy-CH2DCFDA. A representative experiment of 3 independent experiments is shown. B) Jurkat T cells, incubated for 2 h in ROL-free medium, or medium supplemented with 4 µM AR, or 150 µM DHIQ, alone or in combination, were collected and assayed for intracellular levels of ROS by flow cytometry. A representative of 3 independent experiments is shown.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The present studies show that ROL depletion of exogenous sources or blockade of endogenous ROL by AR causes a rapid increase in mitochondria-derived ROS production, mitochondrial depolarization, significant decreases in cellular levels of ATP and NAD+, and caspase-independent cell death in Jurkat T cells and MEFs. A further key finding is the activation of PARP-1, consequent to AR exposure. Using pharmacological inhibitors and PARP-1–/– MEFs, we show that PARP-1 plays a critical role in the cell death execution pathway. These findings highlight the importance of ROL homeostasis in mitochondria and the negative consequences that occur when this balance is upset by ROL deprivation or displacement by AR.

As previously reported, lymphocytes (normal T cells, transformed lymphoblastoid B cells, and T leukemia cells) undergo rapid PCD during nutritional vitamin A deprivation, when internal stores of retinyl-ester become depleted (6 , 16 , 47) . Throughout our work AR is used as a tool to induce an acute state of vitamin A deficiency. AR is a normal metabolite of vitamin A in mammals and insects and is capable of displacing ROL from high-affinity binding sites on serine-threonine kinases (i.e., PKC and cRaf), resulting in diminished responsiveness of these enzymes to the alternate redox activation pathway (11 , 14) . ROL and AR have nearly identical binding affinities, although opposite metabolic effects (11 , 14) . Consequently, the one present in higher concentration in a mixture of the 2 determines the outcome, whether this be mitochondrial integrity, ROS production, ATP synthesis rate, or viability.

Because mammalian cells usually express multiple PKC isoforms, as well as cRaf, each operating in different signal circuits, vitamin A deprivation causes pleotropic cell damage, including depolymerization of the cytoskeleton, loss of osmotic integrity, compromised cell adhesion, and mitochondrial membrane depolarization (8 , 10) . These effects are most readily apparent in growth-factor-deprived cells. Although cumulative damage occurs to multiple organelles, the decisive negative impact leading to programmed cell death (PCD) is likely to be at the mitochondrion, because of increased mitochondrial-derived ROS formation, possibly at complex I or III of the electron transport chain (ETC). At first glance, the action of AR resembles that of fenretinide, which leads to classic apoptosis (48 , 49) . Although oxidative stress, cytochrome c release, and DNA damage are initiated by both, additional hallmarks of apoptosis such as DNA laddering, caspase-3 activation, and externalization of phosphatidylserine to the outer leaflet of the plasma membrane are absent during AR-induced PCD. Instead, AR precipitates an energy crisis that, when not alleviated within an hour, leads to caspase-independent cell death, characterized by rapid loss of plasma membrane integrity. Suppressing ROS production by hypoxia, or dissipating ROS by the antioxidant action of L-NAC or the vitamin E derivative Trolox, effectively counteracts AR-induced PCD.

PARP-1 is an abundant nuclear protein belonging to a large family that includes PARP-2, PARP-3, vault PARP, and tankyrases (50) . Each member of the PARP family shares homology with the C-terminal catalytic domain of PARP-1. PARP-1 is involved in many cellular processes, including DNA single- and double-strand break repair, regulation of transcription, cell cycle, and cell death (21) . In response to DNA damage and other cellular stresses, PARP-1 catalyzes the NAD+-dependent synthesis and attachment of 50–200 ADP-ribose units on {gamma}-carboxyl groups of glutamic acid residues in acceptor proteins such as histones, DNA polymerases, topoisomerases, DNA ligase-2, and several transcription factors (18 , 19) . Recent studies have shown that overactivation of PARP-1 correlates with increased poly(ADP-ribosyl)ation of cellular proteins, rapid utilization and depletion of NAD+ and ATP, and release of death-inducing proteins from the mitochondria, initiating caspase-independent cell death (24 , 26 , 27 , 29) . Inhibition of PARP-1 activity by pharmacological or genetic methodologies is cytoprotective in disease models of ischemia-reperfusion injury, diabetes, inflammatory-mediated injury, ROS-induced injury, and glutamate excitotoxicity (22 , 25 , 51 52 53 54 55) . We have demonstrated that ROL depletion by AR is associated with rapid activation of PARP-1 (within 1 h) and PARP-1-dependent NAD+ and ATP depletion. Conversely, AR-induced cell dysfunction is significantly decreased in the absence of PARP-1. During apoptosis, PARP-1 is often inactivated via cleavage by caspases. However, following AR-treatment of Jurkat cells, PARP-1 is not cleaved (data not shown) and remains functional, probably because the rapid drop in ATP prevents activation of caspases (32 33 34) . The large energy cost associated with PARP-1 action, in terms of NAD+ and ATP consumption, likely contributes to necrotic cell death. Furthermore, our data show that inhibition of PARP-1 does not relieve the initial mitochondrial stress-induced ROS formation. Taken together, our results point to PARP-1 as a key, and thus far unidentified, player in the execution pathway of PCD following ROL depletion.

Our work demonstrates that hypoxia prevents AR-dependent induction of cell dysfunction and death of Jurkat T cells, as well as 143B and bovine pulmonary artery endothelial cells (data not shown). Protection under hypoxia is likely related to decreased mitochondrial-derived ROS and the prevention of ATP and NAD+ depletion. The absence of AR-induced ROS formation under hypoxic conditions suggests a direct action of hypoxia on the mitochondria. Recent studies have shown a close association between HIF-1{alpha} function and mitochondrial respiration. When HIF-1{alpha} is stabilized under hypoxic conditions, mitochondrial complex III-derived ROS furnishes the likely messenger (56 57 58 59) . Up-regulation of pyruvate dehydrogenase kinase 1 (PDK1) (60 , 61) , inactivation of pyruvate dehydrogenase by PDK1, and up-regulation of lactate dehydrogenase (62) are 3 pertinent HIF1-{alpha} mediated events that link AR, ROS, and hypoxia to mitochondrial energy homeostasis. The exploration of these molecules that control the flux of fuel through the ETC may hold the key to identifying the primary interaction partners of ROL in mitochondria and what may go wrong when ROL is either absent or displaced by AR.

In summary, the present study demonstrates that acute ROL deprivation induces a cellular energy crisis and pleomorphic cell dysfunction leading to caspase-independent cell death. This effect can result from decreased mitochondrial ATP production, in addition to ruinous consumption of ATP caused by increased and unabated activation of cellular enzymes and processes utilizing large amounts of ATP. The conceptual advance reported in this study is not that ROL deprivation disables mitochondria, an effect much accentuated by AR, but that ROL acts as positive regulator of cellular bioenergetic processes. ROL, but not AR, is normally available to all tissues and cells. When diminished, cytopathology, or even cell death, ensues. ROL, possibly due to its action on protein kinases, such as PKC and Raf, emerges as a cofactor essential for cellular energy homeostasis.


   ACKNOWLEDGMENTS
 
The authors are indebted to Dr. Valina Dawson (Johns Hopkins University School of Medicine, Baltimore, MD, USA) for sharing PARP-1–/– MEFs. This work was supported by grants from the National Institutes of Health, CA 089362 and DK 069348.

Received for publication April 23, 2008. Accepted for publication July 2, 2008.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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