Modulation of the Protein Kinase Cδ Interaction with the “d” Subunit of F₁F₀-ATP Synthase in Neonatal Cardiac Myocytes

Abstract

The F₁F₀-ATP synthase provides ∼90% of cardiac ATP, yet little is known regarding its regulation under normal or pathological conditions. Previously, we demonstrated that protein kinase Cδ (PKCδ) inhibits F₁F₀ activity via an interaction with the “d” subunit of F₁F₀-ATP synthase (dF₁F₀) in neonatal cardiac myocytes (NCMs) (Nguyen, T., Ogbi, M., and Johnson, J. A. (2008) J. Biol. Chem. 283, 29831–29840). We have now identified a dF₁F₀-derived peptide (NH₂-²AGRKLALKTIDWVSF¹⁶-COOH) that inhibits PKCδ binding to dF₁F₀ in overlay assays. We have also identified a second dF₁F₀-derived peptide (NH₂-¹¹¹RVREYEKQLEKIKNMI¹²⁶-COOH) that facilitates PKCδ binding to dF₁F₀. Incubation of NCMs with versions of these peptides containing HIV-Tat protein transduction and mammalian mitochondrial targeting sequences resulted in their delivery into mitochondria. Preincubation of NCMs, with 10 nm extracellular concentrations of the mitochondrially targeted PKCδ-dF₁F₀ interaction inhibitor, decreased 100 nm 4β-phorbol 12-myristate 13-acetate (4β-PMA)-induced co-immunoprecipitation of PKCδ with dF₁F₀ by 50 ± 15% and abolished the 30 nm 4β-PMA-induced inhibition of F₁F₀-ATPase activity. A scrambled sequence (inactive) peptide, which contained HIV-Tat and mitochondrial targeting sequences, was without effect. In contrast, the cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ facilitator peptide by itself induced the PKCδ-dF₁F₀ co-immunoprecipitation and inhibited F₁F₀-ATPase activity. In in vitro PKC add-back experiments, the PKCδ-F₁F₀ inhibitor blocked PKCδ-mediated inhibition of F₁F₀-ATPase activity, whereas the facilitator induced inhibition. We have developed the first cell-permeable, mitochondrially targeted modulators of the PKCδ-dF₁F₀ interaction in NCMs. These novel peptides will improve our understanding of cardiac F₁F₀ regulation and may have potential as therapeutics to attenuate cardiac injury.

Introduction

The mammalian F₁F₀-ATP synthase is a 16-subunit enzyme complex found in the inner mitochondrial membrane (IM).² It contains an F₁ domain (3α, 3β, γ, δ, and ϵ subunits), which protrudes into the mitochondrial matrix (1–5), and an F₀ domain, which transverses the IM and allows proton entry into the mitochondrial matrix down a concentration gradient. These proton movements produce the driving force for ATP synthesis (4). The F₁ and F₀ domains are connected by a central stalk consisting of the γ, δ, and ϵ subunits and by a peripheral stalk that is made up of the oligomycin-sensitive conferring protein, F6, b, and d subunits (5). The peripheral stalk forms the “stator” that is thought to immobilize the F₁F₀ α and β subunits during ATP synthesis. The central stalk rotates along with the c subunits, which is crucial for proton movement through the F₀ domain and ATP production. The F₁F₀-ATP synthase complex is also regulated by two endogenous inhibitors as follows: inhibitor of F₁ (3, 6–9) and calcium-sensitive binding inhibitor (1, 10). Inhibitor of F₁ binds to the interfaces between α and β subunits to inhibit F₁F₀ activity under conditions of decreased pH and IM potential, such as would occur in ischemia (6–10). The calcium-sensitive binding inhibitor is bound to the enzyme under low mitochondrial Ca²⁺ concentrations and is released from the enzyme as the mitochondrial Ca²⁺ increases (3, 10).

The heart consumes large amounts of ATP, and even though cardiac ATP levels do not change substantially in times of intense activity (due to buffering systems such as creatinine kinase), the F₁F₀ synthase is largely responsible for replenishing the ATP that is used. Despite such modest changes in high energy phosphate levels during strenuous activity, the F₁F₀-ATP synthase is fully capable of responding to increased ATP expenditure. This indicates that in vivo the enzyme is not only regulated by substrate availability but is additionally regulated by biochemical events, such as protein kinase-mediated phosphorylation. There is also evidence that in cardiovascular disease states the F₁F₀-ATP synthase may lose its capacity to increase its activity in response to cellular ATP demands (3). In addition, in chronic cardiomyopathy, ATP levels have been estimated to decline by ∼20% along with an ∼80% drop in phosphocreatinine levels (11), suggesting that chronic energy starvation in the heart may be a major contributor to congestive heart failure (12).

Following severe cardiac ischemia/reperfusion (IR) injury, the type of damage sustained in a heart attack, ATP levels also decline substantially (9, 10). A major component of this drop involves the loss of the mitochondrial IM proton gradient that supplies the energy for F₁F₀-ATP synthase. Therefore, in the early phases of myocardial ischemia, the F₁F₀-ATP synthase becomes inhibited. It then makes a futile attempt to re-establish mitochondrial IM potential by pumping protons out of the mitochondrial matrix. This process requires energy, which is supplied by the F₁F₀ complex, then operating as an ATPase (7, 10). If ischemia is not interrupted, F₁F₀-ATPase activity contributes heavily to the loss of myocardial ATP (7, 10). In addition, the return of aerobic ATP synthesis is impaired following IR injury, but the mechanisms for this delay are not completely known. Under ischemic conditions, the heart attempts to compensate by using glucose as a preferred substrate (instead of predominantly fatty acids) in anaerobic glycolysis. However, anaerobic ATP production is not sufficient to supply the extensive cardiac demands for energy, and it also generates considerable lactic acid, which damages the heart further. The consequent decreases in pH inhibit glycolytic enzymes (7, 10). An accelerated return of aerobic ATP production, following IR injury, would therefore improve the survival of cardiac cells, making the F₁F₀-ATP synthase/ATPase a logical focus of cardiac IR research.

Previous studies have proposed that cardiac pre-conditioning (PC), a cardioprotective response against IR injury, inhibits F₁F₀-ATPase mode activity (13, 14). This could contribute protection by reducing ATP hydrolysis after ischemic insults. There have also been reports that PC induces an improved recovery of cardiac ATP levels following prolonged IR injury (10, 15, 16). This is consistent with the hypothesis that cardiac PC may improve F₁F₀-ATP synthase-mode activity as well. However, to date, the precise roles of the F₁F₀ complex in cardiac IR injury and PC are not fully understood.

Mitochondrial mechanisms involving protein kinase C (PKC) isozymes in cardiac PC and IR injury have received considerable attention (17–21). Identification of the particular isozyme(s) that is involved in these responses is an essential prerequisite for the development of clinically useful pharmacological mimetic agents. In support of this, previous studies have reported a PKCϵ-selective facilitator peptide, and a PKCδ inhibitor peptide can induce cardio-protection from IR injury in cardiomyocytes (22), isolated hearts, and, in vivo, transgenic mice (23–26). The above-mentioned PKCδ inhibitor has also shown great promise in human clinical trials targeting damage from acute myocardial infarction (27). Furthermore, a PKCδ-selective translocation inhibitor, administered immediately following a prolonged ischemic insult, improved the recovery of mouse heart ATP levels during oxygenated reperfusion (25).

In this study, we have identified the first cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ interaction inhibitor and facilitator peptides. We have determined that these peptides modulate the interaction between PKCδ and dF₁F₀ and, consequently, F₁F₀ activity in NCMs. We propose that excessive inhibition of the F₁F₀-ATP synthase by PKCδ may play a significant role in cardiac pathology and that our novel peptides could provide a mechanism for investigating these effects.

EXPERIMENTAL PROCEDURES

Primary NCMs

NCMs were isolated from the hearts of 1-day-old Sprague-Dawley rats as described previously (28, 29). This study was conducted in accordance with Institutional, State, and Federal guidelines for the humane care and use of laboratory animals. Briefly, cells were obtained from hearts by gentle trypsinization at room temperature, and dissociated cells were preplated for 40 min onto 100-mm dishes in medium 199 (M-199) (Invitrogen) containing 10% fetal bovine serum (Hyclone Laboratories). The nonattached cells were plated onto 35- or 100-mm Corning Petri dishes at a density of 800 cells/mm² and incubated at 37 °C in humidified air with 1% CO₂. Myocytes were cultured in M-199 supplemented with vitamin B₁₂ (1.5 mmol/liter), penicillin G (50 units/ml), 0.1 mmol/liter bromodeoxyuridine, and 10% fetal bovine serum through day 4. After the 4th day, cells were then placed in defined M-199 containing 50 units/ml penicillin G, 1.5 mmol/liter vitamin B₁₂, and 10 μg/ml each of transferrin and insulin.

Peptide Synthesis

The 14 sequential dF₁F₀-derived peptides (1 15-mer, 12 16-mers, and 1 18-mer) covering the entire 161-amino acid sequence of dF₁F₀ were synthesized by 21st Century Biochemicals (Marlborough, MA) using Fmoc (N-(9-fluorenyl)methoxycarbonyl) chemistry. Each peptide had a 5-amino acid overlap with the previous peptide. Final sequences of these peptides are shown in Table 1. Several modifications were made to some of the peptides to more closely mimic the physiological forms of the amino acids found in the native dF₁F₀ protein. For example, we did not include the first methionine in dF₁F₀ sequence, as the National Center for Biotechnology Education database indicates that this methionine is not present in the final mature form of dF₁F₀. In addition, we have added N-acetyl and C-carboxyamide groups to the N and C terminus of each peptide, respectively, to properly mimic the peptide bonds normally present in the native protein. Because peptide 14 represents the actual C terminus of the protein, this peptide was made as a free C-terminal carboxyl. Finally, there are several lysines in the native protein (e.g. amino acids 63, 78, 85, 99, 117, and 149) that were incorporated into peptides as N-acetyl-lysines. All peptides were high pressure liquid chromatography-purified (purity ≥90%), and their masses and sequences were verified by collision-induced fragmentation using a Qo-TOF electrospray ionization-mass spectrometer (QSTAR XL Pro, MDX Sciex). All of the peptides were finished as acetate salts, and endotoxin-free water was used for all peptide purification and formulations.

TABLE 1.

Description of sequential peptides derived from the d subunit of adult rat dF₁F₀

Shown are the amino acid sequences of the 14 peptides synthesized as described under “Experimental Procedures” used to determine their effects on PKCδ binding to dF₁F₀. Peptides are listed in order beginning with the N terminus of dF₁F₀ in peptide 1 and ending with the COOH-terminal dF₁F₀ sequence in peptide 14. Each peptide contains a 5-amino acid overlap with the previous sequential peptide. Also note that the N-terminal methionine has been deleted from peptide 1 because it is not thought to be present in the mature dF₁F₀ protein. Other modifications to certain amino acid side chains of our peptides were made to better mimic their in vivo chemistry and are described in detail under “Experimental Procedures.” Shown in boldface superscripts are the actual amino acid numbers corresponding to where each peptide is located in the dF₁F₀ sequence. Amino acids are indicated by universally accepted single letter abbreviations.

Peptide	Amino acid sequence
1	NH2-2AGRKLALKTIDWVSF16-COOH
2	NH2-12DWVSFVEIMPQNQKAI27-COOH
3	NH2-23NQKAIGNALKSWNETF38-COOH
4	NH2-34WNETFHTRLASLSEKP49-COOH
5	NH2-45LSEKPPAIDWAYYRAN60-COOH
6	NH2-56YYRANVDKPGLVDDFK71-COOH
7	NH2-67VDDFKNKYNALKIPVP82-COOH
8	NH2-78KIPVPEDKYTALVDAE93-COOH
9	NH2-89LVDAEEKEDVKNCAQF104-COOH
10	NH2-100NCAQFVTGSQARVREY115-COOH
11	NH2-111RVREYEKQLEKIKNMI126-COOH
12	NH2-122IKNMIPFDQMTIDDLN137-COOH
13	NH2-133IDDLNEVFPETKLDKR148-COOH
14	NH2-144KLDKRKYPYWPHQPIENL161-COOH

Syntheses for cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ interaction inhibitor and facilitator peptides were as follows. Each peptide included either the PKCδ-dF₁F₀ interaction antagonist NH₂-²AGRKLALKTIDWVSF¹⁶-COOH or agonist NH₂-¹¹¹RVREYEKQLEKIKNMI¹²⁶-COOH sequence. An HIV-Tat protein-transducing sequence YGRKKRRQRRR was also placed at the N terminus of each peptide using a cysteine-cysteine linkage as described previously (30). Following the Tat sequence, we included a mitochondrial targeting sequence (MLATRALSLIGKRAISTSVC) derived from the number IV subunit of cytochrome oxidase (31). Finally, a FLAG epitope (DYKDDDDK) was also attached to the C-terminal end of each peptide to monitor their uptake into mitochondria. As a control, a scrambled (inactive) (ADKIGWAVLRTKSLF) version of the PKCδ-dF₁F₀ inhibitor peptide was synthesized with all of the above HIV-Tat, mitochondrial targeting and FLAG domains. The final overall sequence of each of these peptides is shown in Fig. 1A.

FIGURE 1.

Modulation of PKCδ binding to the d subunit of F₁F₀-ATP synthase (dF₁F₀) by dF₁F₀-derived peptides. A, peptides contain the putative PKCδ-dF₁F₀ inhibitor, facilitator, or scrambled (inactive) amino acid sequences, an HIV-Tat protein-transducing (PTD) sequence (YGRKKRRQRRR) for cell uptake, a mitochondrial targeting sequence (MLATRALSLIGKRAISTSVC), and a FLAG epitope (DYKDDDDK). B and C, PKC overlay assays reveal an inhibitor (B) and a facilitator (C) of the PKCδ-dF₁F₀-binding interaction. Purified F₁F₀-ATPase holoenzyme (32) was subjected to SDS-PAGE to resolve individual subunits and transferred onto NCP. NCP strips were “overlaid” with a mixture of purified PKC isozymes (40) in the presence of PKC activators (diacylglycerol and phosphatidylserine) and 0–10 μm concentrations of the putative PKCδ-dF₁F₀ inhibitor (B) or facilitator (C), followed by Western blot analyses using PKCδ-selective antisera (39). Amino acid sequences of each peptide are shown at the top center of B and C. It is important to note that B and C are the only experiments in this study that tested the PKCδ-dF₁F₀ inhibitor and facilitator peptides in the absence of HIV-Tat PTD, mitochondrial targeting, and FLAG domains. Representative autoradiographs are shown, and the histogram values represent mean ± S.E. % of “no peptide control” densitometry values from nine independent experiments for B and six experiments for C. D, cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ inhibitor attenuates PKCδ binding to dF₁F₀. PKC overlay assays were conducted as in B and C in the presence of 0–10 μm concentrations of the cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ inhibitor or scrambled (inactive) peptide. The complete sequences of these peptides are as in A. A representative autoradiograph is shown, and the histogram values represent mean ± S.E. % of densitometry values from six independent experiments. B–D, * indicates statistically significant differences from the no peptide control.

F₁F₀-ATP Synthase Purification

Isolation of the F₁F₀-ATP synthase holoenzyme by chromatography was performed as described previously in detail by Buchanan and Walker (32). Briefly, mitochondria were isolated from five adult Sprague-Dawley rats and were then washed in phosphate buffer and solubilized in buffer containing 20 mm ATP, 20 mm MgSO₄, 0.001% phenylmethylsulfonyl fluoride, 1 mm dithiothreitol, and 1% n-dodecyl-β,d-maltoside. The solubilized mitochondria were precipitated with ammonium sulfate. The ammonium sulfate pellet was resuspended in buffer and dialyzed overnight to remove lipids. The sample was then loaded onto a Q-Sepharose anion exchange column. The F₁F₀-ATP synthase holoenzyme was then eluted using a linear NaCl gradient (from 0 to 1 m), and peak F₁F₀-ATPase fractions were initially identified by the F₁F₀-ATPase activity assay. Subsequently, the presence of the α, β, and d subunits of the F₁F₀-ATPase was confirmed in peak F₁F₀-ATPase activity fractions by Western blots using antisera against each of these subunits. Peak fractions were then pooled and used for overlay binding and other assays. Antisera for the α, β, and d subunits of the F₁F₀-ATPase were obtained from Molecular Probes and Mitosciences and were used in Western blots.

PKC Binding Overlay Assays

The overlay method to detect PKC-binding partners was described previously by Mochly-Rosen and co-workers (33–36). Required activators and co-factors of PKC such as calcium (Ca²⁺), phosphatidylserine (PS, Avanti, Alabaster, AL), and diacylglycerol (DG, Avanti) were included in the assay. Purified F₁F₀ holoenzyme (50 μg) was separated by 13.5% SDS-polyacrylamide gel and transferred onto nitrocellulose paper (NCP). The blots were then cut into strips (0.3 × 5.5 cm) and washed briefly with distilled water. The membrane was then blocked with overlay blocking buffer (50 mm Tris/HCl, pH 7.5, 200 mm NaCl, 3% bovine serum albumin, and 0.1% polyethylene glycol) for 1 h at room temperature. Next, the strips were incubated with overlay buffer containing purified PKC (50 μg/ml) and co-factors for 1 h at room temperature. The strips were washed four times for 5 min with overlay wash buffer. Bound PKC was detected with PKC-isozyme antisera and ECL detection as described below for Western blot analyses.

Cell Lysis and Isolation of Mitochondria from NCMs

100-mm dishes of NCMs were used for mitochondria isolation. The culture medium was removed, and cells were washed twice with Ca²⁺/Mg²⁺-free phosphate-buffered saline containing 1.4 nm calyculin A and 1.44 mm tetrasodium pyrophosphate (NaPP_i). Cells were scraped from the dishes in 400 μl of isotonic MSE buffer (10 mm Tris/HCl, pH 7.5, 220 mm mannitol, 70 mm sucrose, 1 mm EGTA, 0.025% bovine serum albumin, 2 mm taurine, 1.6 mm carnitine, 1.5 mm NaPP_i, 1.5 nm calyculin A, and 10 μg/ml protease inhibitors) containing 100 units/ml trypsin (Sigma) for 20 min on ice. After the incubation period, cells were then subjected to Dounce homogenization. The resulting homogenate was centrifuged twice at 600 × g for 5 min to pellet nuclei and cell debris. Protease inhibitors were added to the final supernatant at 10 μg/ml each, and it was then loaded onto a Percoll/Optiprep gradient as described below (37, 38).

Preparations of Percoll/Optiprep Gradient

The postnuclear supernatant (prepared as described above) was layered over a combination Percoll/Optiprep (Accurate Chemical and Scientific, Westbury, NY) gradient prepared on the same day as follows. Each gradient was prepared in Beckman-Ultraclear 14 × 89-mm centrifuge tubes. The first step in gradient formation involved overlaying 1.74 ml of a 17% (v/v) Optiprep solution on a 1.74-ml cushion of 35% Optiprep solution. Next, 4.35 ml of a 6% (v/v) Percoll solution was layered on top of the 17% Optiprep solution. All Optiprep and Percoll solutions were prepared using MSE buffer as the diluent. Gradients were stored on ice until use. Next, 4.2 ml of postnuclear supernatant was gently layered on top of the 6% Percoll portion of the gradient. All tubes were centrifuged in a Beckman SW.41 swinging bucket rotor at 50,000 × g for 30 min using the lowest acceleration and deceleration speeds. Mitochondria were collected at the 17/35% Optiprep interface from each gradient using a Pasteur pipette and placed on ice until use.

Immunoprecipitation (IP) Experiments

The antisera against the “d” subunit of the F₁F₀-ATPase (dF₁F₀) from Molecular Probes (50 μg) was coupled with Bio-Rad Affi-Gel (1 ml) according to the manufacturer's instructions. The IP was performed as described previously (37, 38) with the addition of 150 mm NaCl to the IP buffer in the three final wash steps.

F₁F₀-ATPase Activity Measurement

The oligomycin-sensitive activity of F₁F₀-ATPase activity was measured using a spectrophotometric assay as described previously (32). Briefly, cells were scraped and lysed by sonication in 400 μl of buffer containing 20 mm Tris/HCl, pH 7.5, 1 mm MgCl₂, 5 mm KCl, and 1 mm EGTA. The F₁F₀-ATPase activity then was measured spectrophotometrically by monitoring the disappearance of NADH, which manifests as a decline in absorbance at 340 nm as NADH is oxidized NAD⁺. The final F₁F₀-ATPase assay buffer contained 25 mm Tris/HCl, pH 7.5, 83 mm sucrose, 4 mm MgCl₂, 25 mm KCl, 1 mm KCN, 1 mm EDTA, 1 mm EGTA, 2 mm ATP, 1.5 mm phosphoenolpyruvate, 5 units of pyruvate kinase, 5 units of lactate dehydrogenase, and 75 μm NADH in a 1-ml cuvette. All assay reagents were obtained from Sigma. The assay was monitored continuously for 1–3 min at 25 °C in the absence and presence of oligomycin (8 μg/ml).

Western Blot Analyses

Western blotting was carried out as described previously (28, 29) using ECL detection (GE Healthcare). Samples were subjected to SDS-PAGE on 12–13.5% acrylamide gels and then transferred onto NCP. The resulting blots were probed for PKC isozymes and F₁F₀ subunits. PKC antisera were obtained from BD Transduction Laboratories or Santa Cruz Biotechnology; F₁F₀ antisera were obtained from Invitrogen, and the FLAG antibody was obtained from Sigma.

Statistical Analyses

Differences between two groups were assessed using unpaired Student's t test, and comparisons among multiple groups were made using one-way analysis of variance with Bonferroni's post hoc test. A p value ≤ 0.05 was considered significant.

RESULTS

Design of dF₁F₀-derived Peptides

Our previous studies in NCMs demonstrated an interaction between PKCδ and the d subunit of F₁F₀-ATP synthase (dF₁F₀), which correlated with 4β-PMA- and hypoxia-induced inhibition of F₁F₀-ATPase activity (39). We next surveyed which amino acid sequence(s) within the dF₁F₀ protein was responsible for its binding to PKCδ. Currently, there are no commercially available recombinant sources of mammalian dF₁F₀ protein. For this reason, we synthesized 14 sequential peptides (Table 1) covering the entire amino acid sequence of dF₁F₀ as described under “Experimental Procedures.”

PKC Overlay Assays Reveal an Inhibitor and Facilitator of the PKCδ dF₁F Interaction

We have previously demonstrated that PKCδ, but not PKCα, -ϵ, or -ζ, selectively binds to the dF₁F₀ protein in overlay assays (39). In Fig. 1, B and C, purified F₁F₀-ATPase holoenzyme was isolated from adult rat heart mitochondria by chromatography (32). Next, individual F₁F₀-ATPase subunits were resolved by SDS-PAGE and then transferred to NCP. The resulting NCP blots were incubated with a mixture of purified rat brain PKC isozymes (40) and PKC activators, in the presence of increasing concentrations of each of the 14 dF₁F₀-derived peptides shown in Table 1. For brevity, data for only the two peptides with the most optimal effects on the PKCδ-F₁F₀ binding interaction are shown (Fig. 1, B and C). PKCδ binding was detected by anti-PKCδ antisera and ECL techniques as described previously (39). It should be noted that four different F₁F₀-ATPase preparations, three different PKC preparations, and two different lots of peptides were used to obtain the data shown in Fig. 1, B–D. A major finding of our study was that one of the peptides (NH₂-²AGRKLALKTIDWVSF¹⁶-COOH) demonstrated a dose-dependent inhibition of the PKCδ-dF₁F₀ binding interaction (Fig. 1B). When 0.3 μm concentrations of this peptide were included in the overlay assay, we observed a 37 ± 9% (n = 9) inhibition of PKCδ binding to dF₁F₀. The calculated IC₅₀ for this peptide was 0.216 μm. NCBI database searches revealed that the amino acid sequence of this inhibitor is not found in any known proteins except the mammalian F₁F₀ d subunit.

A second major finding of our study was that one of the remaining peptides (NH₂-¹¹¹RVREYEKQLEKIKNMI¹²⁶-COOH) facilitated the binding of PKCδ to dF₁F₀ in the overlay assay (Fig. 1C). In the presence of PKC activators, 0.3 μm concentrations of this peptide potentiated PKCδ binding to dF₁F₀ by 229 ± 76% (n = 6). The EC₅₀ for this peptide was 0.71 μm. These results suggested that we had identified two peptides that modulate PKCδ binding to dF₁F₀ in opposite directions; one peptide was an inhibitor and the second peptide acted as a facilitator of this interaction.

Addition of HIV-Tat, Mitochondrial Targeting, and FLAG Sequences Do Not Reduce the Efficacy of the PKCδ-dF₁F₀ Inhibitor in Overlay Assays

In vitro binding of PKCδ to dF₁F₀ in the presence of increasing concentrations of the HIV-Tat/mitochondrial targeting domain-containing PKCδ-dF₁F₀ inhibitor (Fig. 1A) was monitored using the approach outlined in Fig. 1, B and C. When this inhibitor peptide (100 nm) was included in overlay assays, we observed a 42 ± 14% (n = 6) reduction in PKCδ binding to dF₁F₀ (Fig. 1D). The inactive version of this peptide had no effect on the binding of PKCδ to dF₁F₀ (Fig. 1D). Therefore, the modified antagonist/inhibitor exhibited a dose-dependent inhibition of the PKCδ-dF₁F₀ binding interaction (Fig. 1D), which appeared to be modestly more potent (IC₅₀ = 0.144 μm) than the inhibitor sequence alone (Fig. 1B, IC₅₀ = 0.216 μm).

Incubation of NCMs with the Cell-permeable, Mitochondrially Targeted PKCδ-dF₁F₀ Inhibitor and Facilitator Peptides Results in Their Delivery into Mitochondria

We next incubated NCMs in the presence or absence of the mitochondrially targeted PKCδ-dF₁F₀ facilitator (Fig. 2A) or inhibitor (Fig. 2B) peptides (100 nm) for 2 h at 37 °C. Extracellular media were then removed, and cells were washed twice with phosphate-buffered saline. Next, each group of cells was homogenized, and mitochondria were isolated using differential centrifugation and Percoll/Optiprep density gradients as described previously (38, 39). In Fig. 2, lane 1, 0.5 μg of the FLAG epitope-tagged facilitator (Fig. 2A) or inhibitor (Fig. 2B) peptide alone was subjected to SDS-PAGE to confirm that we could use anti-FLAG antisera in Western blots to detect our peptides. As shown, each peptide was readily detected. In Fig. 2, A and B, lane 2 is similar to lane 1 except each peptide was first incubated with trypsin (100 units/ml) for 20 min on ice prior to conducting SDS-PAGE and Western blots with anti-FLAG antisera. Note the substantial loss in FLAG immunoreactivity confirming that each peptide was sensitive to trypsin digestion. In Fig. 2, A and B, lanes 3 and 4, we subjected mitochondria isolated from NCMs that were not incubated with any peptide to SDS-PAGE and Western blot analyses with anti-FLAG antisera. As predicted, no FLAG immunoreactivity was observed in these samples. In contrast, Fig. 2, A and B, lanes 5 and 6 represent samples prepared by isolating density gradient-purified mitochondria from NCMs that were preincubated for 2 h with 100 nm concentrations of either the PKCδ-dF₁F₀ facilitator (Fig. 2A) or inhibitor (Fig. 2B) peptides. Note the strong anti-FLAG immunoreactivity observed in Fig. 2, A and B, lane 5. These results indicate that extracellular treatment of NCMs with our peptides resulted in substantial mitochondrial uptake of each peptide. To confirm that the peptides were not simply adhering to extramitochondrial surfaces, we treated the mitochondria with 100 units/ml trypsin for 20 min on ice (Fig. 2, A and B, lane 6). Note that this treatment did not diminish the anti-FLAG immunoreactivity (Fig. 2A, lane 5 versus 6), despite the extreme sensitivity of our peptides to digestion by trypsin (Fig. 2, A and B, lane 1 versus 2). These results suggested that our FLAG-tagged peptides were not likely to be simply adhering to extramitochondrial surfaces and that they do enter NCM mitochondria.

FIGURE 2.

Exposure of neonatal rat cardiac myocytes to the HIV-Tat-coupled, mitochondrially targeted versions of the PKCδ-dF₁F₀ facilitator or inhibitor leads to uptake into gradient-purified mitochondria. PKCδ-dF₁F₀ facilitator (A, lanes 1 and 2) or PKCδ-dF₁F₀ inhibitor (B, lanes 1 and 2) were incubated in the absence (lane 1) or presence (lane 2) of 100 units/ml trypsin for 20 min at 4 °C then subjected to SDS-PAGE, followed by Western blot analysis using anti-FLAG antisera. NCMs were also treated in the absence (A, lanes 3 and 4) or presence (A, lanes 5 and 6) of 100 nm concentrations of the PKCδ-dF₁F₀ facilitator or in the absence (B, lanes 3 and 4) or presence (B, lanes 5 and 6) of the PKCδ-dF₁F₀ inhibitor for 2 h. Mitochondria (Mito.) were then purified as under “Experimental Procedures.” Isolated mitochondria were then subjected to SDS-PAGE and Western blot analysis using antisera directed against the FLAG epitope. These mitochondria were incubated in the absence (lane 5) and presence (lane 6) of 100 units/ml trypsin for 20 min at 4 °C immediately after isolation. Representative autoradiographs for FLAG immunoreactivity are shown, and histograms represent mean ± S.E. % of maximal densitometry values from three independent experiments.

Modulation of the 4β-PMA-induced Inhibition of F₁F₀-ATPase Activity by Cell-permeable, Mitochondrially Targeted PKCδ-dF₁F₀ Inhibitor and Facilitator Peptides in NCMs

Cells were preincubated with the cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ antagonist/inhibitor or agonist/facilitator peptides (10 nm) for 2 h at 37 °C followed by a 30 nm 4β-PMA treatment for 30 min. Cells were processed and mitochondrial F₁F₀-ATPase activity assayed as described previously (39). 30 nm 4β-PMA treatment inhibited the basal, oligomycin-sensitive, F₁F₀-ATPase activity by 41 ± 2.6% (n = 13) (Fig. 3, leftmost set of bars). Incubation of NCMs with the scrambled sequence inactive control peptide (Fig. 1A) for 2 h did not alter the 4β-PMA-induced inhibition of F₁F₀-ATPase activity (Fig. 3, 2nd set of bars from left). In contrast, preincubation of NCMs with 10 nm concentrations of the PKCδ-dF₁F₀ inhibitor peptide completely abolished the 4β-PMA-induced inhibition of F₁F₀-ATPase activity (Fig. 3, 3rd set of bars from the left).

FIGURE 3.

Incubation of NCMs with the cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ peptides modulate 4β-PMA-induced inhibition of F₁F₀-ATPase activity. NCMs were treated with no peptide (left) or 10 nm concentrations of scrambled (inactive), inhibitor, or facilitator peptides (Fig. 1) for 2 h prior to 30 nm 4α- or 4β-PMA for 30 min. The oligomycin-sensitive F₁F₀-ATPase activity was measured spectrophotometrically (“Experimental Procedures”). Results are expressed as mean ± S.E. from eight independent experiments, each conducted in triplicate using samples from eight different myocyte preparations. Asterisks indicate statistically significant differences from no peptide group.

The effects of the mitochondrially targeted PKCδ-dF₁F₀ facilitator peptide was somewhat predictable in that it promotes PKCδ binding to dF₁F₀ by itself in overlay assays (Fig. 1C). We hypothesized that because the interaction of PKCδ with dF₁F₀ induces PKCδ-mediated inhibition of F₁F₀-ATPase activity that the PKCδ-dF₁F₀ facilitator peptide, administered by itself, should inhibit F₁F₀-ATPase activity. In fact, this turned out to be the case as is shown in Fig. 3, last set of bars. Interestingly, the combination of the PKCδ-dF₁F₀ facilitator with 4β-PMA did not induce synergistic inhibition of F₁F₀-ATPase activity. We believe that this is because at the concentrations of 4β-PMA and PKCδ-dF₁F₀ facilitator peptide used, we observe maximal co-IP of PKCδ with dF₁F₀ (Fig. 4B), which is to say that treatment of NCMs with either 4β-PMA or the PKCδ-dF₁F₀ facilitator peptide induces maximal PKCδ co-IP with dF₁F₀.

FIGURE 4.

Extracellular treatment of NCMs with the cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ interaction of modulatory peptides attenuates 4β-PMA-induced co-immunoprecipitation of PKCδ with dF₁F₀. A, NCMs were preincubated in the absence of peptide (lanes 1 and 2) or in the presence of 100 nm concentrations of the scrambled sequence inactive peptide (lane 3), the PKCδ-dF₁F₀ inhibitor (lane 4), or the PKCδ-dF₁F₀ facilitator (lane 5) for 2 h. The complete amino acid sequences of these peptides are as in Fig. 1A. Next, all groups, except lane 1, were exposed to 100 nm 4β-PMA for 30 min. Mitochondria were then isolated as in Fig. 2. Mitochondrial proteins were subjected to immunoprecipitation reactions using dF₁F₀ antisera coupled to Affi-Gel. The resulting IPs were probed for PKCδ in Western blot analyses. As is shown, only the cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ inhibitor peptide attenuated 4β-PMA-induced co-IP of PKCδ with dF₁F₀ antisera (lane 4). A typical autoradiograph is shown at the top of the figure. Bars in the histogram represent mean ± S.E. densitometry values taken from five independent experiments each conducted on mitochondria isolated from a separate myocyte preparation. * indicates statistically different from co-IP values obtained with 4β-PMA alone in the absence of any peptide (lane 2). Con, control. B, extracellular treatment of NCMs with the cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ facilitator peptide induces the PKCδ-dF₁F₀ co-IP. NCMs were administered no treatments (lane 1) or treatments with 30 nm 4β-PMA for 30 min (lane 2) and with 1 (lane 3) or 10 (lane 4) nm extracellular concentrations of the PKCδ-dF₁F₀ interaction facilitator peptide (Fig. 1A) for 2 h (lanes 3 and 4). Mitochondria were isolated; co-IP reactions were carried out, and Western blots were probed for PKCδ as in Fig. 4A. Under basal conditions there is little co-IP between dF₁F₀ and PKCδ (lane 1). In contrast, 4β-PMA treatment causes a dramatic increase in the co-IP of PKCδ with dF₁F₀ (lane 2). Lanes 3 and 4 are co-IPs taken from NCM mitochondria isolated from cells that were treated with 1–10 nm concentrations of the PKCδ-dF₁F₀ facilitator in the complete absence of 4β-PMA. Note the dose-dependent induction of the PKCδ-dF₁F₀ co-IP by the facilitator peptide alone. The autoradiograph shown is from a typical experiment. Histogram bars represent mean ± S.E. densitometry values expressed as % of maximal from three independent experiments each conducted on a separate myocyte preparation. * indicates statistically different from control co-IP values. # indicates statistically different from co-IP values obtained for 4β-PMA alone in the absence of any peptide.

Treatment of NCMs with the Cell-permeable, Mitochondrially Targeted PKCδ-dF₁F₀ Inhibitor Attenuates the 4β-PMA-induced Co-IP of PKCδ with dF₁F₀

In the experiments represented in Fig. 4A, NCMs were incubated for 2 h at 37 °C in the absence or presence of our mitochondrially targeted peptides (Fig. 1A) followed by a 30-min exposure to 100 nm 4β-PMA. Cells were next homogenized, and mitochondria were isolated by differential centrifugation and Percoll/Optiprep density gradients (38). Mitochondria were then lysed and subjected to IP experiments using dF₁F₀ antisera chemically coupled to Affi-Gel. IPs were subjected to SDS-PAGE and Western blot analyses using anti-PKCδ antisera. Consistent with our previous study (39), 4β-PMA treatment of NCMs induced the co-IP of PKCδ with dF₁F₀ (Fig. 4A, lane 2). This co-IP is preferential for the PKCδ isozyme as we have previously shown that phorbol ester treatment of NCMs does not induce co-IP of PKCα,- ϵ, or -ζ isozymes with dF₁F₀ (39). When cells were incubated in the presence of the mitochondrially targeted versions of the inactive scrambled sequence or the PKCδ-dF₁F₀ facilitator peptides (100 nm), there was no effect on the 4β-PMA-induced PKCδ-dF₁F₀ co-IP (Fig. 4A, lane 2 versus lanes 3 and 5). However, the cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ inhibitor attenuated the 4β-PMA-induced co-IP of PKCδ with dF₁F₀ antisera by 50 ± 15% (n = 4) (Fig. 4A, lane 4). These results are consistent with the hypothesis that the PKCδ-dF₁F₀ inhibitor diminishes the 4β-PMA-induced inhibition of F₁F₀-ATPase activity (Fig. 3) by attenuating the PKCδ-dF₁F₀ interaction in NCMs.

Treatment of NCMs with the Cell-permeable, Mitochondrially Targeted PKCδ-dF₁F₀ Facilitator Peptide Induces the PKCδ-dF₁F₀ Co-IP

To determine the effects of the PKCδ-dF₁F₀ facilitator peptide (10 nm) (Fig. 1A) on the PKCδ co-IP with dF₁F₀, we conducted the experiments represented in Fig. 4B. We initially hypothesized that this peptide would enhance this 4β-PMA-induced effect. However, we did not observe any such enhancement (Fig. 4A). It is possible that at these concentrations, each stimulus could induce the PKCδ-dF₁F₀ co-IP maximally, and therefore we did not observe synergy when the two compounds were applied together. Fig. 4B supports the conclusion that treatment of NCMs with either 4β-PMA (lane 1 versus 2) or the PKCδ-dF₁F₀ facilitator (lane 1 versus 3 and 4) alone induces the PKCδ-dF₁F₀ co-IP. Furthermore, the effects of the facilitator peptide were dose-dependent as 10 nm concentrations of the PKCδ-dF₁F₀ facilitator increased the PKCδ-dF₁F₀ co-IP by 2.1 ± 0.1-fold (n = 3) greater than 1 nm concentrations of the peptide (Fig. 4B). These results are consistent with the PKCδ-dF₁F₀ facilitator inhibiting F₁F₀-ATPase activity (Fig. 3) by induction of the PKCδ-dF₁F₀ interaction in NCMs (Fig. 4B).

PKCδ-dF₁F₀ Inhibitor Attenuates and the PKCδ-dF₁F₀ Facilitator Induces Inhibition of F₁F₀-ATPase Activity in Vitro

We have previously demonstrated that co-incubation of recombinant PKCδ with chromatographically purified F₁F₀-ATPase from rat heart mitochondria inhibits F₁F₀-ATPase activity (39). This inhibition occurs in the absence of PKC activators and is amplified in their presence (39). We next determined if our HIV-Tat-coupled, mitochondrial targeted PKCδ-dF₁F₀ interaction modulators (Fig. 1A) could inhibit or facilitate PKCδ-mediated inhibition of F₁F₀-ATPase activity in vitro (Fig. 5). Addition of recombinant PKCδ to purified F₁F₀-ATPase in the absence of PKC activators leads to a 1.7 ± 0.1-fold inhibition of F₁F₀-ATPase activity. In the presence of the PKC activators DG and phosphatidylserine (PS), PKCδ inhibited F₁F₀ activity by 92 ± 5% (Fig. 5). The scrambled sequence, inactive control peptide had no impact on these PKCδ-mediated responses. In contrast, 10 nm concentrations of the cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ inhibitor attenuated the PKCδ-mediated inhibition of F₁F₀-ATPase in the absence of PKC activators by 28 ± 3% (n = 3) (Fig. 5, lanes 2 versus 3). A much more dramatic effect of the PKCδ-dF₁F₀ inhibitor peptide was observed in the presence of PKC activators as it reduced recombinant PKCδ-induced inhibition of F₁F₀-ATPase activity in the presence of DG/PS by 7.6 ± 0.6-fold (n = 3) (Fig. 5, lanes 5 versus 6).

FIGURE 5.

Cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ inhibitor and facilitator peptides have opposing effects on PKCδ-mediated inhibition of F₁F₀-ATPase activity in vitro. Rat heart F₁F₀-ATPase was purified from isolated mitochondria using the methods of Buchanan and Walker (32) and recombinant PKCδ, purified from an Sf9 cell expression system, was obtained from BIOSOURCE. F₁F₀-ATPase (50 μg) activity was monitored spectrophotometrically in the absence (lane 1) or presence (lanes 2–7) of 1.5 μg of recombinant PKCδ. Assays were conducted in the absence (lanes 1–4) or presence (lanes 5–7) of the PKC activators DG (3.2 μg/ml) and PS (240 μg/ml). The effects of 10 nm concentrations of the cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ interaction inhibitor (lanes 3 and 5) or facilitator (lanes 4 and 7) on PKCδ-mediated inhibition of F₁F₀-ATPase activity were then evaluated. Note that the PKCδ-dF₁F₀ inhibitor attenuated PKCδ-mediated inhibition of F₁F₀-ATPase activity whether or not DG/PS was present in the assay. In contrast, the facilitator modestly enhanced PKCδ-mediated inhibition of F₁F₀-ATPase in the absence of DG/PS. Results shown are mean ± S.E. results from four independent experiments. * indicates statistically different from control (F₁F₀ only). # indicates statistically different from F₁F₀ + recombinant PKCδ group.

We next determined whether inclusion of the cell-permeable, mitochondrially targeted PKCδ-dF₁F₀ facilitator in the PKCδ/F₁F₀-ATPase add-back assay amplified PKCδ inhibitory actions. Consistent with this, in the absence of DG/PS, we found 55 ± 13% (n = 3) greater PKCδ-mediated inhibition of F₁F₀-ATPase when 10 nm concentrations of the PKCδ-dF₁F₀ facilitator peptide were included in the assay (Fig. 5, compare lanes 2 and 4 to lane 1). Furthermore, the PKCδ-dF₁F₀ facilitator peptide modestly enhanced PKCδ-mediated inhibition of F₁F₀ activity when the PKC activators DG/PS were present in the assay (Fig. 5, lanes 5 versus 7). Collectively, these data support the hypothesis that both DG/PS and the PKCδ-dF₁F₀ facilitator peptide promote the PKCδ interaction with dF₁F₀ to inhibit F₁F₀ activity.

DISCUSSION

The human heart consumes up to 20 times its mass in ATP each day, and F₁F₀-ATP synthase may be the most crucial enzyme for meeting these intense energy requirements. Therefore, a better understanding of the regulation of the F₁F₀ complex under normal and pathological states is an extremely important goal. Unfortunately, very little has been learned over the past 2 decades regarding cardiac F₁F₀ function. Recently, our laboratory identified a novel and potent inhibitory regulation of this enzyme complex by the PKCδ isozyme. The response involves PKCδ binding to the d subunit of the F₁F₀-ATP synthase (dF₁F₀) and is induced by phorbol ester and hypoxia in NCMs (39).

The role of mitochondrial PKCδ in cardiac PC (41–43) and IR injury (44–47) is controversial. Our studies support the notion that whether PKCδ mediates cardiac protection or injury may be context-dependent. For example, it would be beneficial if PKCδ inhibited ATP hydrolysis during prolonged ischemia. In support of this, numerous studies have reported that cardiac PC inhibits F₁F₀-ATPase activity (13, 14). Alternatively, PKCδ could exacerbate injury if it inhibits F₁F₀-ATP synthase-mode activity during early reperfusion after a prolonged ischemic insult. It is well known that one characteristic of cardiac IR injury involves a delayed recovery of aerobic ATP synthesis (10, 44). We hypothesize that such a delay in the recovery of aerobic ATP production could contribute heavily to cardiac myocyte cell death. Consequently, disrupting the PKCδ-dF₁F₀ interaction at the time of cardiac reperfusion could facilitate a more rapid return of aerobic ATP production following IR injury. We therefore developed novel tools to pursue the regulation of the F₁F₀ complex by PKCδ interaction with dF₁F₀ in live heart cells.

There are no published reports for the expression of mammalian F₁F₀-ATP synthase subunits in bacteria, baculovirus/Sf9 systems. The first phase of this study was therefore to prepare a sequential series of peptides derived from the 161-amino acid dF₁F₀ protein (Table 1). We then used these peptides in PKC overlay assays to determine which ones could antagonize the in vitro binding of PKCδ to dF₁F₀. We reasoned that such a peptide would act as a competitive antagonist of PKCδ binding to dF₁F₀. The peptide shown in Fig. 1B fulfilled these criteria and was selected for the development of a compound that could disrupt the PKCδ-dF₁F₀ interaction in NCMs.

An unanticipated finding was that the dF₁F₀-derived peptide NH₂-¹¹¹RVREYEKQLEKIKNMI¹²⁶-COOH caused a dose-dependent potentiation of PKCδ binding to dF₁F₀ in overlay assays (Fig. 1C). The mechanism by which this occurs is at present unknown, but there have been reports of peptides derived from individual PKC isozymes that can enhance their binding to target proteins and facilitate their intracellular trafficking (47, 48). For example, peptide sequences derived from individual PKC isozymes called pseudo-RACK sequences have been reported to accomplish this (34). In fact, expression of pseudo-RACK sequences for the PKCϵ and -δ isozymes into mouse hearts in vivo produces cardioprotection and exacerbates damage (50), respectively, following a severe IR challenge. Complete characterization of the mechanisms of the PKCδ-dF₁F₀ facilitator peptide will require further study and is outside of the scope of the current work. However, we are very intrigued by this molecule because it not only has effects in in vitro PKC overlay assays (Fig. 1C), it also inhibits F₁F₀-ATPase activity (Fig. 3) and induces the PKCδ-dF₁F₀ co-IP in NCMs (Fig. 4B).

The next phase of our study involved application of a technology that would allow high efficiency delivery of our PKCδ-dF₁F₀ inhibitor and facilitator peptides into NCM mitochondria to determine whether they could modulate F₁F₀-ATPase activity. It has been previously reported by Dowdy and co-workers (51) that intraperitoneal injection of rodents with the β-galactosidase protein chemically coupled to the HIV-Tat protein transduction domain sequence, used in this study, allowed high efficiency delivery of the protein to virtually all tissues in the body of the animal. Our peptides presented an additional challenge in that the F₁F₀-ATP synthase exists in the mitochondrial IM. Therefore, to enhance or disrupt PKCδ interaction with dF₁F₀, our modulatory peptides would not only need to enter cells, they would also need to be delivered to the IM where F₁F₀-ATPase exists. Of interest, the HIV-Tat protein transduction domain has previously been reported to carry the apoptosis repressor with caspase recruitment domain (ARC) protein into NCM mitochondria when the fusion protein was applied extracellularly (52). The ARC protein is ∼40 kDa in size, which is considerably larger than any of the peptides we tested in this study. However, we used a cysteine-cysteine chemical linkage (30, 50) to attach the HIV-Tat protein transduction domain to our peptides, which supposedly allows the Tat protein transduction domain sequence to be cleaved off once the peptide enters the cell. We therefore felt it was necessary to include a mitochondrial targeting sequence (31) to our peptides.

It is well known that ∼99% of mitochondrial proteins are encoded by nuclear genes and are translated in the cytosol before being transported into mitochondria. All proteins taken up by mitochondria are thought to have the mitochondrial targeting amino acid sequences located, for the most part, on their N terminus (53). This sequence binds to various chaperones and an elaborate array of mitochondrial protein carriers located in the outer and inner mitochondrial membranes (53). The mitochondrial targeting sequence we selected is one of the best studied and is derived from the number IV subunit of cytochrome oxidase (31). Our results indicate high efficiency mitochondrial uptake of our inhibitor, facilitator, and scrambled sequence peptides when NCMs are treated extracellularly with them (Fig. 4). In addition to their detection in mitochondria using FLAG immunoreactivity in Western blots, we found our peptides able to achieve sustainable levels in mitochondria suggesting they were not being completely proteolyzed or otherwise lost. In support of this, FLAG immunoreactivity in Western blots co-migrated with pure peptide alone and did not change in molecular weight (Fig. 2, A and B, lanes 1 versus 5). In addition, the cell-permeable inhibitor and facilitator peptides altered F₁F₀-ATPase activity and the PKCδ-dF₁F₀ co-IP (Figs. 3–5) in opposing directions, whereas the scrambled sequence control peptide had no effect.

There have been reports of nonspecific effects of HIV-Tat protein transduction sequences when introduced into cells (49). In general, toxicities were observed using much higher concentrations (20–500 μm) of the HIV-Tat protein transduction domain peptide than we used. In addition, our peptides were prepared as acetate salts (rather than the generally more toxic trifluoroacetate counter-ions) and using endotoxin-free water, which may also reduce nonspecific toxicities. Therefore, we do not believe that our peptides caused nonspecific actions in these studies. In support of this, our cell-permeable, mitochondrial targeted PKCδ-dF₁F₀ inhibitor had similar effects to attenuate PKCδ binding to dF₁F₀ in in vitro overlay assays (Fig. 1D). We also developed a control peptide that contains all of the HIV-Tat, mitochondrially targeting, and FLAG domains as well as a scrambled amino acid sequence, derived from the inhibitor peptide (Fig. 1A). This scrambled sequence peptide was taken up into mitochondria, but it failed to modify basal or 4β-PMA-induced changes in F₁F₀-ATPase activity (Fig. 3) or co-IPs (Fig. 4A). Finally, using Live/Dead vital staining (Invitrogen) and cardiac troponin I release, we observed no cytotoxicity following a 2-h incubation with up to 1 μm extracellular concentrations of the peptides (data not shown).

Collectively, these results provide the first proof-of-principle evidence in support of the opposing mechanisms of the PKCδ-dF₁F₀ inhibitor and facilitator peptides. We propose that these novel molecules will allow us to either attenuate or induce PKCδ-mediated inhibitory effects on the F₁F₀ complex to assess the role(s) of this interaction in different cardiac responses. We are currently focusing on the utility of these novel peptides for reducing injury and improving energetics in adult rat models of IR injury in our laboratory.