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The Journal of Immunology, 2007, 178: 869-876.
Copyright © 2007 by The American Association of Immunologists, Inc.

HIV-1 Trans Activator of Transcription Protein Elicits Mitochondrial Hyperpolarization and Respiratory Deficit, with Dysregulation of Complex IV and Nicotinamide Adenine Dinucleotide Homeostasis in Cortical Neurons1

John P. Norman2,*,{dagger}, Seth W. Perry*, Karl A. Kasischke, David J. Volsky|| and Harris A. Gelbard*,{ddagger},§

* Department of Neurology (Child Neurology Division), Center for Aging and Developmental Biology, University of Rochester School of Medicine and Dentistry, Rochester, NY 14642; {dagger} Department of Environmental Medicine, University of Rochester School of Medicine and Dentistry, Rochester, NY 14642; {ddagger} Department of Pediatrics, University of Rochester School of Medicine and Dentistry, Rochester, NY 14642; § Department of Microbiology and Immunology, University of Rochester School of Medicine and Dentistry, Rochester, NY 14642; Department of Neurosurgery, Center for Aging and Developmental Biology, University of Rochester School of Medicine and Dentistry, Rochester, NY 14642; and || Molecular Virology Division, St. Luke’s-Roosevelt Hospital Center and College of Physicians and Surgeons, Columbia University, New York, NY 10019


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
HIV-1 causes a common, progressive neurological disorder known as HIV-associated dementia (HAD). The prevalence of this disorder has increased despite the use of highly active antiretroviral therapy, and its underlying pathogenesis remains poorly understood. However, evidence suggests that some aspects of HAD may be reversible. To model the reversible aspects of HAD, we have used the HIV-1 neurotoxin trans activator of transcription protein (Tat) to investigate nonlethal changes in cultured neurons. Exposure of rodent cortical neurons to sublethal concentrations of Tat elicits mitochondrial hyperpolarization. In this study, we used the cationic lipophilic dye rhodamine 123 to confirm this observation, and then performed follow-up studies to examine the mechanism involved. In intact neurons, we found Tat elicited a rapid drop in internal mitochondrial pH, and addition of Tat to purified mitochondrial extracts inhibited complex IV of the electron transport chain. To correlate enzyme activity in mitochondrial extracts with results in intact cells, we measured neuronal respiration following Tat exposure. Cortical neurons demonstrated decreased respiration upon Tat treatment, consistent with inhibition of complex IV. We examined mitochondrial Ca2+ homeostasis using a mitochondrial targeted enhanced yellow fluorescent protein-calmodulin construct. We detected a decrease in mitochondrial calcium concentration following exposure to Tat. Finally, we measured the energy intermediate NAD(P)H after Tat treatment, and found a 20% decrease in the autofluorescence. Based on these findings, we suggest that decreased NAD(P)H and calcium concentration contribute to subsequent respiratory decline after exposure to Tat, with detrimental effects on neuronal signaling.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Human immunodeficiency virus-associated dementia (HAD)3 is a progressive neurological disorder that results in motor, cognitive, and social dysfunction. Although the phenotype and severity of HAD have changed in the last decade, largely due to highly active antiretroviral therapy, the neuroradiologic, neuropsychologic, and epidemiologic studies suggest that HAD prevalence has in fact increased since 2000. Thus, long-lasting structural and functional changes to the CNS, as a result of HIV type 1 (HIV-1) infection, continue to be a healthcare problem of global significance (1).

Because administration of highly active antiretroviral therapy can temporarily ameliorate symptoms of HAD, it is speculated that there is a reversible component of HAD due to a metabolic encephalopathy (2). Because HIV-1 causes neurologic impairment without directly infecting neurons, we and others have chosen to model the effects of HIV-1 on neurons by studying the actively released HIV-1 protein, trans activator of transcription protein (Tat), and its effects on neuronal physiology as an initial step toward understanding what happens to neuronal bioenergetics under conditions that lead to synaptic dysfunction (3).

The trans activator of transcription protein, or Tat, is actively secreted by infected cells such as T cells and macrophages/microglia (4), thereby causing effects on bystander cells such as neurons (5, 6, 7). These effects may be mediated either at the cell surface, or following intracellular uptake of Tat via its arginine-rich basic domain (8, 9). Acute neurotoxic effects of Tat may therefore occur through several mechanisms, including overactivation of N-methyl-D-asparate (NMDA) receptors to cause excitotoxicity and apoptosis (10, 11, 12, 13, 14, 15). Furthermore, Tat exposure also results in the release of neurotoxic effector molecules from macrophages/microglia and neurons themselves, including NO, TNF-{alpha}, and platelet-activating factor (16, 17, 18). Finally, Tat can move into mitochondrial spaces via its basic domain (also known as a protein transduction domain (PTD)) (19).

Mitochondria are responsible for energy metabolism, ion homeostasis, and apoptosis regulation in eukaryotic cells (20, 21). Mitochondria produce energy by transferring electrons through a chain of progressively reduced protein complexes embedded in the inner mitochondria membrane, while using the resulting energy to drive H+ against the concentration gradient, into the inner-membrane space (out of the mitochondrial cytoplasm) (22). The resulting H+ gradient flows back though the F1/F0 complex (into the mitochondria), where ADP and Pi are condensed into ATP. This entire complex is referred to as the electron transport chain (ETC). The electrochemical gradient generated by the ETC is referred to as the proton motive force ({Delta}p). The {Delta}p is comprised of two forces, the mitochondrial membrane potential ({Delta}{psi}m) (charge gradient) and the mitochondria pH (pHm) gradient (concentration gradient). This relationship is represented by the equation {Delta}p = {Delta}{psi}m + pHm. The approximate physiological values are –220 mV = –180 mV + –40 mV (interior negative) (23). Under normal physiologic conditions, {Delta}p remains relatively constant, but in disease states mitochondria may fail to maintain the {Delta}p, leading to altered mitochondrial energetics (24). In neurons, one of the most frequently reported events leading to a loss in {Delta}p (depolarization) is a rapid change in intracellular ion homeostasis, particularly from Ca2+ (25, 26, 27).

Our lab has reported previously that addition of HIV-1 Tat to cortical neurons hyperpolarizes {Delta}{psi}m (28); mitochondrial hyperpolarization has also been reported in other models of HIV-1 infection (29) as well as in other model systems (30, 31, 32). Although hyperpolarization has been demonstrated to be detrimental to mitochondrial function and cell survival, it remains uncertain whether it signifies a pathophysiologic mechanism for HIV-1-induced neurodegeneration. In this current study, we systematically investigated the mitochondrial targets through which Tat might act to cause hyperpolarization. Our results suggest a dominant effect of changes in Ca2+ homeostasis and in the decrease in NAD(P)H.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Reagents

rHIV-1 Tat1–72 and biologically inactive {Delta}31–61 Tat were generously provided by A. Nath (University of Kentucky, Lexington, KY) (33). The gp120 (IIIB) was obtain from the AIDS Research and Reference Reagent Program. YC3.1 constructs (34, 35) were a generous gift from the laboratory of N. Demaurex (Department of Physiology, University of Geneva, Geneva, Switzerland) and were originally developed by R. Tsien (Department of Pharmacology, University of California, San Diego, CA). The dyes rhodamine 123 (rhod123), SNARF-1, and MitoTracker Green were purchased from Molecular Probes. Neurobasal medium and B27 supplement (with and without antioxidants) were purchased from Invitrogen Life Technologies. All other chemicals and reagents were purchased from Sigma-Aldrich.

Primary neuronal cell culture

Primary neuronal cortical cultures were harvested and prepared from embryonic day 18 Sprague-Dawley rat pups with modifications to the protocol described by Brewer et al. (36). In brief, the cortices were isolated from a litter of E18 rats, and the meninges and extraneous tissue were removed. The cortices were incubated in 2 ml of Ca2+/Mg2+-free HBSS (HBSS with 10 mM HEPES (pH 7.3)) with gentamicin (50 µg/ml) and 0.25% trypsin for 15 min at 37°C. The cells were centrifuged at 1000 rpm for 5 min, washed twice with HBSS (with Ca2+/Mg2+), then dissociated in Neurobasal medium supplemented with glutamate, gentamicin, and B27 supplement (Invitrogen Life Technologies) by 10 passages through a 0.9-mm bore pipette tip. Dissociated cells were counted using the trypan blue viability assay and were plated on poly(D-lysine)-coated cell culture plastic and incubated in a humidified atmosphere of 5% CO2/95% air at 37°C. The supplemented Neurobasal medium is modified for an antioxidant-free culture, as described by Perry et al. (29), and inhibits the growth of glial cell populations. The resulting cultures are 98% pure neuronal cultures (36). Cultures were used for experiments at days in vitro 11–14, unless otherwise noted.

Image capture and image analysis

The use of fluorescent dyes allowed us to image physiological changes in real time. The following setup was used for all experiments in which a fluorophore was used. Samples were placed on a DC60 warming stage (Linkam Scientific Instruments) and maintained at 37°C for the duration of the experiment. A single field was monitored during the course of the experiment, and images were taken using an Olympus IX-70 microscope with x40 objective and an Apogee KX32ME charge-coupled device camera. Images were analyzed using Scanalytics IPLab software. Quantification of the neuronal fluorescent intensity was determined by the total sum of the recorded pixel values within the specified region of interest, for each image series captured.

Isolated mitochondrial proteins

Cortical neuronal mitochondria were prepared from P17 Sprague-Dawley rat pups, as described by Maciel et al. (37), with minor modifications. Rats were decapitated, and their brains were rapidly removed and placed into 5 ml of ice-cold isolation buffer (295 mM mannitol, 65 mM sucrose, 2 mM HEPES, 2 mg/ml BSA, and 1 mM EGTA (pH 7.4)). The cortices were dissected, placed in 2 ml of isolation buffer, and manually homogenized using a Wheaton 10-ml glass/Teflon mortar and pestle on ice. The resulting homogenate was centrifuged at 4°C for 5 min at 1,500 x g in a Beckman S4180 rotor (Beckman Coulter). The supernatant was subsequently centrifuged at 12,000 x g for 8 min at 4°C. The supernatant was discarded, and the pellet was suspended in 10 ml of isolation buffer. Protein concentrations were determined using the Bradford protein assay (Bio-Rad). Mitochondrial protein was stored at –80°C until use in ETC complex assays, except for complex II/III-linked assay, which was used immediately after protein isolation.

Activities of the mitochondrial ETC complexes in rat brain

The activities of the ETC complexes I, II, II/III, and IV were measured using isolated brain mitochondria, as previously described (38), with several modifications. Briefly, 50 µg of mitochondrial protein was freeze thawed for use in the measurement of complex I and II activity. Complex I activity was measured by monitoring NADH oxidation at 340-nm absorption. To determine NADH reduction not due to complex I, 2 µg/ml rotenone was added, and the results were subtracted from the initial results to give the complex I oxidation of NADH. Complex II activity was monitored through the reduction in absorption of dichlorophenolindophenol at 600 nm. To assess maximum activity, mitochondrial protein was incubated in the presence of 20 mM succinate for 10 min before the experiment. Complex III activity was indirectly measured by a linked assay between complexes II and III (succinate-hexacyanoferrate reductase assay). Activity is determined by the decrease in absorbance as the hexacyanoferrate (III) at 420 nm. In order for complex II to be maximally activated, mitochondrial protein (200 µg) was incubated in the presence of 20 mM succinate for 10 min before the experiment. Complex IV activity was assessed using the cytochrome c oxidase assay kit (Sigma-Aldrich), as per the manufacturer’s instructions. The activity of complex IV was monitored by the oxidation of cytochrome c at 550 nm by mitochondrial protein (10 µg). To determine the background levels of cytochrome c oxidation, 100 µM potassium cyanide was added to the mitochondrial protein and the results were subtracted to obtain the final complex IV activity.

Polarography

Oxygen consumption by cortical neuronal cultures was monitored using a Clark-type electrode (Hansatech Instruments, PP System). Briefly, cortical neurons were cultured on glass coverslips (7 x 20 mm) at a density of 105 cells/coverslip. A slotted Teflon peg was inserted into the chamber into which three coverslips were placed (Fig. 4B). To determine cellular respiration, cortical neurons were placed into Neurobasal medium containing the experimental treatment, and the recording chamber was sealed. Respiration was monitored for 10 min. The complex IV inhibitor, potassium cyanide, was used to subtract background from the recordings.


Figure 4
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  		FIGURE 4. HIV-1 Tat decreases cortical neuron respiration. A, Cortical neuronal (105 neurons/coverslip, 3 coverslips per experiment) respiration was examined to determine the effect of HIV Tat on complex IV in a physiological system. Treatment with Tat elicited a concentration-dependent decrease in neuronal respiration. Addition of {Delta}31–61 Tat had no effect on cortical neuronal respiration. The viral envelope gp120 (IIIB) was also studied to demonstrate the specificity of Tat’s effects on complex IV respiration. gp120 alone had no effect on complex IV respiration. B, The chamber schematic that was used to measure neuronal respiration is illustrated here. A Teflon peg with slots carved into it was placed above the oxygen electrode, and there was space on either side of the peg to allow free diffusion of drug/oxygen. Coverslips (three) were placed into the slots, and the chamber was filled with Neurobasal medium containing the proper treatment. The chamber was sealed, and measurements were recorded for 10 min. The mean oxygen consumption was plotted, and treated neurons were compared with control for statistical significance (n = 4).

 
Assessing mitochondrial {Delta}{psi}m

The {Delta}{psi}m was determined, as described (23, 39), with several slight modifications. Cortical neuronal cultures were incubated with 10 µM rhod123 under normal incubation conditions for 30 min. The rhod123-containing medium was removed, and the cultures were placed in CO2-insensitive, pH-stable Leibovitz’s L-15 medium (Invitrogen Life Technologies) and incubated at 37°C in room air for 15 min to allow for the rhod123 fluorescent signal to reach a steady state. Culture treatment was applied directly to the bath medium. Samples were imaged using a Texas Red filter (Chroma Technology) at 560/645 (ex/em). The high concentration of rhod123 causes the fluorophore to quench, which inversely correlates to the {Delta}{psi}m.

Measuring mitochondrial pH

Assessment of mitochondrial pH was performed, as described (40). Briefly, cortical neurons were incubated with 10 µM SNARF-1 for 10 min under normal incubation conditions. The SNARF-1 was washed out, and the cultures were gently washed twice with Neurobasal (without antioxidants), then placed in the incubator for 4 h to allow the SNARF-1 dye to unload from the cytoplasm, while being retained by the mitochondria (40). Cortical neurons were placed in Leibovitz’s L-15 medium (Life Technologies) and imaged using a Texas Red filter (Chroma Technology) at 560/645 (ex/em). Only fluorescent areas with a distinct punctate pattern (pixel area >2, but <200) were quantified to ensure mitochondrial pH rather than residual cytosolic pH. The appropriate treatment was added directly to the incubating medium. Calibration was accomplished by incubating cortical neurons at varying pHs and loading with SNARF-1, as above. Nigericin (10 µM), an H+/K+ exchanger, was added to the neurons and incubated for 10 min. SNARF-1 fluorescence was quantified at the various pH values, and a pH dose curve was generated and used in subsequent experiments.

Internal mitochondria Ca2+ measurements

Mitochondrial Ca2+ measurements were accomplished through the use of a mitochondrial targeted enhanced yellow fluorescent protein (EYFP)-calmodulin construct. Briefly, cortical neurons were transfected using Lipofectamine 2000 (Invitrogen Life Technologies), as per the manufacturer’s instructions (2 µl of lipofectamine to 1 µg of DNA). Cortical neurons were incubated at 5% CO2/95% air at 37°C for 24 h before the lipofectamine was removed and returned to the incubator for 4 days. Transfected cultures were placed in Leibovitz’s L-15 medium (Life Technologies) and incubated at 37°C in room air for 25 min before imaging. The appropriate experimental treatment was added directly to the medium and monitored the fluorescence resonance energy transfer from cyan fluorescent protein (CFP) at 425/480 (ex/em) (custom filter; Chroma Technology) to EYFP fluorescence at 425/540 (ex/em) (Lucifer Yellow filter; Chroma Technology) (34, 41). Data are expressed as the ratio between the emission of CFP to EYFP at 480–540 nm.

Two-photon microscopy of NAD(P)H fluorescence

Two-photon NAD(P)H fluorescence imaging was performed using techniques as described by Kasischke et al. (42). The two-photon imaging setup consisted of an upright Olympus Fluoview FV300 laser-scanning microscope equipped with external Hamatsu Photomultiplier tubes. For two-photon excitation of NAD(P)H, a Coherent Chameleon femtosecond Ti:Sa laser system was tuned to 740 nm. Cortical neurons were plated at a density of 106/35 mm2 dish, and baseline NAD(P)H fluorescence images were obtained before the addition of Tat. Dynamics in the cellular NAD(P)H after addition were imaged using several 10-min time series with an interval of 60 s between image acquisition. Image analysis was performed with ImageJ (National Institutes of Health).

Data and statistical analysis

Raw data were pooled from several independent experiments and expressed as mean ± SEM, with error bars indicating SEM. Where applicable, the percentage of control and percentage of control SEM for each treatment condition were calculated by dividing the raw means and raw SEMs by the control condition raw mean. Data were compared by unpaired Student’s t test using a two-tailed distribution and unequal variance. A probability of p < 0.05 was considered statistically significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Acute exposure to Tat increases mitochondrial {Delta}{psi}m in cortical neurons

The measurement of the {Delta}{psi}m is used to gauge mitochondrial functional integrity (24). Using tetramethylrhodamine ethyl and methyl esters in nonquenching mode, our laboratory has demonstrated previously that Tat elicits mitochondrial hyperpolarization in a biphasic manner (28). To further validate the observation that acute Tat exposure induces mitochondrial hyperpolarization in cortical neurons, the cationic, lipophilic fluorescent dye rhod123 was used in quenching mode, as an alternate means of measuring {Delta}{psi}m. The cationic nature of rhod123 allows for its selective accumulation in the negatively charged matrix of mitochondria, resulting in a quantitative measurement of the {Delta}{psi}m (23). The autoquenching properties of rhod123 at high concentrations result in an inverse correlation between dye concentration and fluorescent signal, meaning a decrease in signal reflects higher retention of rhod123 and a rise in (i.e., hyperpolarization of) {Delta}{psi}m (23). Conversely, an increase in the rhod123 signal is associated with an unloading or unquenching event, suggesting a depolarization of the {Delta}{psi}m. Our controls confirmed this expected behavior (see below).

By this method, acute exposure to Tat was shown to elicit a hyperpolarization of the {Delta}{psi}m. Rhod123 staining exhibited the expected mitochondrial labeling, delineated by a distinct punctate staining pattern. A moderate concentration, 1 µg/ml (~80 nM) Tat, caused a 10% decrease in rhod123 fluorescence, indicating increased rhod123 partitioning into the mitochondria due to increased {Delta}{psi}m (Fig. 1A). The decrease in fluorescence occurred rapidly following Tat exposure, indicating a possible direct interaction with the mitochondria. We have shown previously that Tat-induced changes in rhodamine dye concentrations in mitochondria are not a consequence of alterations in plasma membrane potential or mitochondrial morphology (28).


Figure 1
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  		FIGURE 1. Tat induces mitochondrial hyperpolarization. A, Treatment with 1 µg/ml Tat (n = 6; T = Tat; O = oligomycin; F = FCCP) resulted in a 10% loss in rhod123 fluorescence, indicating a rise in the {Delta}{psi}m (*, p < 0.005). Addition of oligomycin at t = 18 elicited a further decrease in rhod123, as was expected with the inhibition of the F0/F1 complex. Addition of FCCP caused a rapid increase in rhod123 fluorescence, indicating a rapid depolarization of the mitochondria. B, Treatment with 1 µg/ml {Delta}31–61 Tat resulted in no significant change in rhod123 fluorescence (n = 3; DT = {Delta}31–61 Tat; O = oligomycin; F = FCCP). C, Cortical neurons treated with oligomycin (n = 4; O = oligomycin; F = FCCP) to demonstrate that the rhod123 is functioning correctly, as reflected by a decrease in fluorescence following oligomycin exposure (indicating a hyperpolarization of the mitochondria) (*, p < 0.01; #, p < 0.05). The mean values (percentage of control) of rhod123 fluorescent were plotted, and treated time points were compared with control time points for statistical significance.

 
Addition of the F0/F1-ATPase inhibitor oligomycin at a late time point (20 min) following Tat treatment induced a further hyperpolarization (Fig. 1A) as the oligomycin inhibited H+ movement into the mitochondria. Addition of the mitochondrial specific protonophore, carbonyl cyanide-4(trifluoromethoxy)phenylhydrazone (FCCP), caused a rapid loss of {Delta}{psi}m, as indicated by the rise (i.e., unquenching) of mitochondrial rhod123 fluorescence (Fig. 1A). The inactive Tat mutant ({Delta}31–61) was used as a negative control, due to its inability to elicit mitochondrial hyperpolarization, its inability to activate NMDA receptors, and its lack of penetrance through biological membranes (28, 43). The {Delta}31–61 Tat mutant had no effect on {Delta}{psi}m (Fig. 1B), demonstrating that full-length, active Tat is required to induce mitochondrial hyperpolarization in intact cells. Finally, as a control, cells were exposed to oligomycin alone; as expected, this elicited a loss in rhod123 fluorescence (hyperpolarization) (Fig. 1C).

A decrease in mitochondrial pH in response to acute Tat exposure

Because {Delta}{psi}m is only one component of the {Delta}p, we also measured neuronal pHm in response to Tat treatment. Under physiological conditions, the forces {Delta}{psi}m and pHm combine to establish {Delta}p at ~–220 mV (23). Using the protocol for selective mitochondrial targeting, the pH-sensitive dye SNARF-1 allows for imaging pHm (40). In cellular systems, SNARF-1 fluorescence increases with increasing pH; the change in fluorescence can be used to generate a standard pH curve and thereby reveal the pH of the SNARF-1-loaded mitochondria.

SNARF-1 mitochondrial targeting is accomplished by loading neuronal cultures with the dye, washing the excess out of the culture plates, and then allowing SNARF-1 to distribute into the cytoplasm (40). This technique takes advantage of anionic transporters located in the plasma membrane that unload SNARF-1 from the cytoplasm. The mitochondria are deficient in these transporters, and, as a result, after several hours of unloading, SNARF-1 produces distinct punctae in neurons that correlate to mitochondria. These punctae were then quantified to establish mitochondrial pH (Fig. 2, A–C, example puncta shown by arrows). SNARF-1 fluorescence decreased rapidly after Tat application, indicating a decrease in pHm (Fig. 2D). Application of Tat (1 µg/ml) decreased intramitochondrial pH from pH 7.6 to 7.3, reflecting the increased H+ in the mitochondrial interior (Fig. 2D). As expected, the addition of the protonophore FCCP caused a rapid decrease in pHm (i.e., influx of H+) (Fig. 2D), whereas oligomycin, which inhibits the F0/F1 complex (thereby inhibiting H+ transfer to the interior of the mitochondria), elicited a slight increase in pHm (from pH 7.55 to 7.75), also as expected (Fig. 2F). The biologically inactive {Delta}31–61 Tat mutant (1 µg/ml) did not elicit a change in pHm (Fig. 2E).


Figure 2
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  		FIGURE 2. Effects of Tat on pHm. A, A representative field (x300 final magnification) of control rodent cortical neurons mitochondria selectively loaded with SNARF-1. B, The same field stained with MitoTracker Green. C, Overlay of fields A and B to demonstrate SNARF-1’s mitochondrial specific staining. The resulting punctae (examples delineated by arrows) were quantified to determine the pHm. D, Neurons treated with 1 µg/ml Tat (n = 5; T = Tat; O = oligomycin; F = FCCP) resulted in a decrease (from 7.6 to 7.3) in pHm (*, p < 0.005; #, p < 0.01; +, p < 0.05). E, Treatment with the biologically inactive {Delta}31–61 Tat (1 µg/ml) (n = 3; DT = {Delta}31–61 Tat; O = oligomycin; F = FCCP) resulted in no significant change in pH (*, p < 0.05). F, Inhibition of F0/F1 complex (inhibition of H+ flow into the mitochondria) with oligomycin (n = 3; O = oligomycin; F = FCCP) resulted in a rise in the pHm (*, p < 0.005). The mean values (percentage of control) of SNARF-1 fluorescent were plotted, and treated time points were compared with control time points for statistical significance.

 
ETC activity

Next, we investigated the role of individual ETC complexes in mediating mitochondrial hyperpolarization induced by Tat. To do this, purified mitochondrial extracts were exposed to Tat, and ETC activity was measured; the results are depicted in Fig. 3A. Complexes I and II showed no significant change in activity when Tat (1 µg/ml) or {Delta}31–61 Tat mutant (1 µg/ml) was added. Full-length Tat caused a 29% decrease in the complex II/III-linked activity compared with control. This assay uses complex II as the electron donor to complex III, which transfers the electron to the complex III artificial substrate (hexacyanoferrate III), whose change in absorption is measured. Because there was no observed change in complex II, we can infer that the change in complex II/III activity can be solely attributed to complex III. The {Delta}31–61 Tat mutant control showed no change in complex III activity, suggesting that the PTD is responsible for the observed decrease in complex III activity.


Figure 3
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  		FIGURE 3. Tat inhibits mitochondrial complexes III and IV. A, Isolated mitochondrial protein was used to measure individual mitochondrial complex activity (n = 4 for each complex). Tat treatment (1 µg/ml) had no effect on complex I and II activity, whereas complexes III and IV showed a significant decrease. Treatment with {Delta}31–61 Tat (1 µg/ml) had the surprising effect of decreasing complex IV activity (*, p < 0.001; #, p < 0.005; +, p < 0.05). B, Examination of Tat’s effect on complex IV (n = 3) revealed a concentration-dependent inhibition of function (*, p < 0.05).

 
Complex IV demonstrated the most dramatic decrease in activity, with a reduction to 49% of control upon addition of 1 µg/ml Tat. The {Delta}31–61 Tat mutant also elicited a striking decrease in complex IV activity (to 60% of control levels, when added at 1 µg/ml). This suggests that regions outside the PTD may be, at least in part, responsible for the effects on complex IV. To further test this hypothesis, we used another HIV-1 protein, gp120 (IIIB), to demonstrate the specificity of Tat’s effect. We examined complex IV activity in the presence of 10 ng/ml and 1 µg/ml gp120 (IIIB) and observed no significant difference in activity (107.4 + 10.6 and 109.3 + 12.1% control, respectively; n = 5).

Dose-response studies of complexes III and IV revealed no further inhibition of complex III with higher Tat concentrations (data not shown), but did reveal a concentration-dependent inhibition of complex IV (Fig. 3B). A concentration of 5 ng/ml, 50 ng/ml, and 10 µg/ml inhibited complex IV activity to 84, 47, and 35% of control, respectively (Fig. 3B). In aggregate, the data can be modeled by a sigmoidal dose-response curve, suggesting saturability of complex IV inhibition by Tat.

Measuring the respiration of cortical neurons

To evaluate the potential significance of Tat’s effects on mitochondrial bioenergetics, we performed experiments using whole cells. ETC complex activity is difficult to measure directly in an intact cellular system. However, complex IV is responsible for the majority of oxygen consumption in cellular systems, and therefore, we quantified oxygen consumption using a Clark-type electrode as a surrogate measure of the activity of complex IV (44, 45).

Cortical neurons demonstrated a concentration-dependent decrease in oxygen consumption following Tat treatment (Fig. 4A). A threshold concentration of 100 ng/ml Tat significantly inhibited respiration to 64% of control in cortical neurons, and a higher dose (1 µg/ml) further inhibited oxygen consumption to 47% of control. Although {Delta}31–61 mutant Tat decreased complex IV activity in mitochondrial extracts (Fig. 3B), treatment of intact neurons with this Tat mutant did not show any significant effect on respiration (Fig. 4A). This most likely reflects the fact that {Delta}31–61 Tat does not cross biological membranes efficiently, because it lacks the PTD that is required for intracellular uptake. Thus, it has no effect in intact cells. To demonstrate the specificity of Tat’s effect on respiration, the HIV envelope gp120 was also used as a control (Fig. 4A). gp120 alone did not cause a significant decrease in respiration in cortical neurons.

Effects of HIV Tat on NAD(P)H fluorescence

NADH is used as a substrate for complex I in mitochondria, donating two electrons to be transported in the ETC. The resulting NAD+ is then reduced by the Krebs cycle enzymes malate-dehydrogenase, succinate dehydrogenase, and citrates synthase to regenerate NADH (46).

It has been shown that the mitochondrial NAD(P)H, but not ADP or Pi, is a likely candidate as a regulator of mitochondrial respiration in vivo (47). Imaging intrinsic NAD(P)H fluorescence changes with two-photon microscopy provides direct visualization of the effect of HIV Tat on mitochondrial metabolism in cortical neurons. Through the use of two-photon microscopy, we monitored a significant, linear decrease (17%) in mitochondrial fluorescence of NAD(P)H within ~10 min (Fig. 5A) after application of HIV-1 Tat. The effect was specific for Tat, because the application of {Delta}31–61 Tat did not result in an observable fluorescence change. As expected, rotenone increased the NAD(P)H fluorescence to ~145% of control by preventing complex I from oxidizing NADH to NAD+ (Fig. 5B).


Figure 5
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  		FIGURE 5. Effects of Tat on NAD(P)H fluorescence in cortical neurons. A, Two-photon microscopy was used to examine the effect of Tat (1 µg/ml) and {Delta}31–61 Tat (1 µg/ml) on NAD(P)H fluorescence (n = 5) in cortical neurons (*, p < 0.05; #, p < 0.005). B (inset), Application of rotenone (10 µM) caused a 40% rise in the NAD(P)H fluorescence (*, p < 0.05; #, p < 0.005).

 
Measuring mitochondrial calcium changes due to Tat exposure

We next turned our attention to whether Tat affected mitochondrial stores of Ca2+. To do this, we transfected cortical neurons with an EYFP-calmodulin construct specifically targeted to the mitochondria via the cytochrome oxidase subunit VIII targeting sequence, and then quantified the FRET between the EYFP to CFP fluorescence as a measure of Ca2+ concentration (34).

Transfected cortical neurons demonstrated a punctate fluorescent labeling confined to mitochondria (Fig. 6A, examples of mitochondria punctae demonstrated by arrows). When compared with untreated and {Delta}31–61 Tat controls, (1 µg/ml) Tat induced a rapid decrease in overall CFP:EYFP fluorescence (Fig. 6B). This decrease in calmodulin fluorescence indicates a decrease in free (unbound) mitochondrial Ca2+, and this loss of ionic charge may result in the observed changes in {Delta}{psi}m. The application of {Delta}31–61 mutant Tat did not elicit a change in Ca2+, which is in agreement with the fact that this Tat mutant also failed to alter {Delta}{psi}m in intact neurons.


Figure 6
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  		FIGURE 6. Mitochondrial calcium response to HIV-1 Tat treatment. A, Representative image of cortical neuron transfected with the mitochondrial targeted EYFP-calmodulin (x400 final magnification), which fluoresces in a punctate pattern demonstrating proper mitochondrial targeting (examples delineated by arrows). B, Application of 1 µg/ml Tat (n = 4; TX = treatment) to transfected neurons initiated a loss of CFP:EYFP calmodulin fluorescence, indicating a loss in mitochondrial calcium (*, p < 0.05; #, p < 0.001). Application of the positive control, ionomycin (2 µM) and calcium (10 mM), increased the CFP:EYFP calmodulin fluorescence.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Mitochondrial hyperpolarization in neurons has only recently been reported (30, 48, 49), and previous studies related to mitochondrial hyperpolarization in the context of HIV-1 infection have focused on T cells (29). We have reported previously Tat-induced mitochondrial hyperpolarization in cortical neurons (28), and in this study we investigated possible mechanisms underlying this phenomenon.

The first question we sought to answer involved determining which ETC component is responsible for the hyperpolarization. Tat application had the unexpected effect of causing acidification of the interior of the mitochondria - that is to say, an increased H+ (Fig. 2D). Taken by itself, this decrease in mitochondrial pH should cause the mitochondrial membrane to depolarize rather than hyperpolarize, which raises a conceptual conundrum. On the one hand, total mitochondrial negativity ({Delta}{psi}m) (as assessed by cationic dyes exhibiting their expected physicochemical behavior), increases with Tat treatment (Fig. 1A and Perry et al.) (29). (Ample evidence suggests Tat is not simply facilitating entry of these cationic dyes into mitochondria: 1) Tat is cationic and should not exhibit charge-interactions with these dyes; 2) mutant ({Delta}31–61), heat-inactivated, and preabsorbed Tat do not confer the same effects; 3) Tat washout (before dye incubation in nonquenching mode) does not prevent observed hyperpolarization; and 4) Tat’s downstream mediators, platelet-activating factor and TNF-{alpha}, also cause increased {Delta}{psi}m (28) (data not shown).) In contrast, the pHm data indicate that there is an increase in H+ ions inside Tat-treated mitochondria (Fig. 2D), and that Tat inhibits complex IV (Figs. 3A and 4A). Taken separately, these latter two pieces of data would suggest that the mitochondria should be depolarizing rather than hyperpolarizing. However, pHm is not an independent variable; thus, we had to examine the individual complexes of the ETC.

Complexes I and II did not show any changes in activity in response to Tat (Fig. 3A), but complexes III and IV showed significant decreases following Tat exposure (Fig. 3A). Complex IV appeared to be the most sensitive, as inhibition was observed with a dose as low as 5 ng/ml Tat (Fig. 3B). These results with mitochondrial extracts were confirmed by the observed decline in cellular respiration in intact neurons exposed to HIV-1 Tat (Fig. 4A). Importantly, the {Delta}31–61 Tat mutant (which lacks the PTD necessary to effectively penetrate the plasma membrane (50)) and HIV-1 gp120 had no effect on cellular respiration, confirming the specificity of Tat’s effects. The decrease in respiration is in accordance with the observed decrease in pHm; as complex IV becomes inhibited, it loses the ability to transfer H+ from the interior of the mitochondria to the outside. This result, however, does not agree with the observed hyperpolarization, and so we next examine the role of NAD(P)H in cortical neurons.

In our analysis of NAD(P)H in Tat-treated neurons, we observed a brisk decrease to 83% of control following Tat exposure (Fig. 5), as would be consistent with respiratory inhibition and intramitochondrial proton accumulation (decreased pH) from reduced ETC activity. We speculated that the TCA cycle could be inhibited by Tat, and thus, NADH levels would begin to decrease as NADH is oxidized to NAD+ by complex I without replenishment of the reduced NADH by the TCA cycle. Future studies beyond the scope of this study will focus on dissecting the causes for the decrease in NADP(P)H. To shed light on the apparent contradictory observations between {Delta}{psi}m and pHm, we analyzed mitochondrial Ca2+ homeostasis following Tat exposure. Application of Tat induced a decrease in free (unbound) mitochondrial Ca2+ vs untreated and mutant Tat controls (Fig. 6B). This loss of Ca2+ would be expected to have consequences that would include neurite retraction, degeneration of synaptic transmission, and malfunction of other Ca2+-related cellular signals (28). In addition to having deleterious effects on neuronal signaling, the loss in free mitochondrial Ca2+ could result in the observed hyperpolarization of mitochondrial membranes. A loss of the Ca2+ ionic charge would increase the total anionic charge of the mitochondria, thus increasing {Delta}{psi}m. The loss of intramitochondrial Ca2+ ionic charge is one explanation that may resolve the observed paradox of increase mitochondrial H+ concentration (i.e., lower pHm) in the face of observed mitochondrial hyperpolarization.

Interestingly, when 50 ng/ml Tat was applied to isolated complex IV protein (Fig. 3B), there was an approximate loss of 50% of associated activity. To observe a similar reduction in complex IV activity in intact cells, a dose of 1 µg/ml Tat was required (Fig. 4A). Thus, a 20-fold greater concentration of Tat concentration was required to elicit the same effect in intact neurons vs isolated mitochondria. This suggests that ~5% of bath-applied Tat can interact directly with cortical mitochondria within intact neurons. This is the first estimate of cellular penetrance of Tat using a physiological measurement that we are aware of. It also has significant implications for dendrites in proximity to an HIV-1-infected macrophage or astrocyte, as we now know a measurable portion of the released Tat from these cells is able to affect the function of mitochondria in adjacent neurons.

Our observations lead us to speculate that TCA cycle activity may be compromised following Tat exposure, in intact cells. A decrease in TCA activity may account for the decrease in NAD(P)H fluorescence. Our findings further suggest that the observed decrease in mitochondrial calcium may also play a pivotal role in mitochondrial membrane hyperpolarization due to the loss of the positive charge associated with free calcium. The loss of mitochondrial calcium also may reflect a decreased buffering capacity of the mitochondria. This has important implications as Tat, at higher concentrations, can also activate NMDA receptors and the effects of subsequent calcium influx could amplify glutamate-induced excitotoxicity.

Thus, we have demonstrated several previously unreported metabolic effects of the HIV-1 neurotoxin Tat, including complex IV inhibition, a decrease in free mitochondrial Ca2+, and alterations in the mitochondrial redox state reflected by the oxidation of NAD(P)H. Together, these findings provide a conceptual framework for understanding how the HIV-1 protein Tat might negatively impact mitochondrial energetics, robbing neurons of luxury functions such as synaptic transmission. This in turn provides us with potential new molecular and cellular therapeutic targets for treatment of the neurologic symptoms of HIV-1 infection.


   Acknowledgments
 
We acknowledge Dr. Stephen Dewhurst for critically reading this manuscript, and Dr. Maiken Nedergaard for the generous use of the two-photon equipment.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by the Geoffrey Waasdorp Pediatric Neurology Fund and National Institutes of Health Grants PO1MH64570, R01MH56838, and R01MH071176 (to H.A.G.); T32ES07026 (to J.P.N.); and PO1NS31492 (to D.J.V. and H.A.G.). This work was also supported by the American Heart Association Grant 0635595T (to K.A.K.) and by the ALS Association Grant 1112 (to K.A.K.). Back

2 Address correspondence and reprint requests to John P. Norman, Center for Aging and Developmental Biology, University of Rochester School of Medicine and Dentistry, 601 Elmwood Avenue, Box 645, Rochester, NY 14642. E-mail address: John_Norman{at}urmc.rochester.edu Back

3 Abbreviations used in this paper: HAD, HIV-1-associated dementia; {Delta}{psi}m, mitochondrial membrane potential; {Delta}p, proton motive force; ETC, electron transport chain; FCCP, carbonyl cyanide-4(trifluoromethoxy)phenylhydrazone; NMDA, N-methyl-D-asparate; pHm, mitochondrial pH; PTD, protein transduction domain; rhod123, rhodamine 123; Tat, trans activator of transcription protein; EYFP, enhanced yellow fluorescent protein; CFP, cyan fluorescent protein; NAD(P)H, signifies both NADH and NADPH. Back

Received for publication August 16, 2006. Accepted for publication October 27, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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