Mitochondrial Alterations Induced by the p13II Protein of Human T-cell Leukemia Virus Type 1
2002; Elsevier BV; Volume: 277; Issue: 37 Linguagem: Inglês
10.1074/jbc.m203023200
ISSN1083-351X
AutoresDonna M. D’Agostino, Laura Ranzato, Giorgio Arrigoni, Ilaria Cavallari, Francesca Belleudi, Maria Rosaria Torrisi, Micol Silic‐Benussi, Tiziana Ferro, Valeria Petronilli, Oriano Marin, Luigi Chieco‐Bianchi, Paolo Bernardi, Vincenzo Ciminale,
Tópico(s)Endoplasmic Reticulum Stress and Disease
ResumoHuman T-cell leukemia virus type 1 encodes a number of "accessory" proteins of unclear function; one of these proteins, p13II, is targeted to mitochondria and disrupts mitochondrial morphology. The present study was undertaken to unravel the function of p13II through (i) determination of its submitochondrial localization and sequences required to alter mitochondrial morphology and (ii) an assessment of the biophysical and biological properties of synthetic peptides spanning residues 9–41 (p139–41), which include the amphipathic mitochondrial-targeting sequence of the protein. p139–41 folded into an α helix in micellar environments. Fractionation and immunogold labeling indicated that full-length p13II accumulates in the inner mitochondrial membrane. p139–41 induced energy-dependent swelling of isolated mitochondria by increasing inner membrane permeability to small cations (Na+, K+) and released Ca2+ from Ca2+-preloaded mitochondria. These effects as well as the ability of full-length p13II to alter mitochondrial morphology in cells required the presence of four arginines, forming the charged face of the targeting signal. The mitochondrial effects of p139–41 were insensitive to cyclosporin A, suggesting that full-length p13II might alter mitochondrial permeability through a permeability transition pore-independent mechanism, thus distinguishing it from the mitochondrial proteins Vpr and X of human immunodeficiency virus type 1 and hepatitis B virus, respectively. Human T-cell leukemia virus type 1 encodes a number of "accessory" proteins of unclear function; one of these proteins, p13II, is targeted to mitochondria and disrupts mitochondrial morphology. The present study was undertaken to unravel the function of p13II through (i) determination of its submitochondrial localization and sequences required to alter mitochondrial morphology and (ii) an assessment of the biophysical and biological properties of synthetic peptides spanning residues 9–41 (p139–41), which include the amphipathic mitochondrial-targeting sequence of the protein. p139–41 folded into an α helix in micellar environments. Fractionation and immunogold labeling indicated that full-length p13II accumulates in the inner mitochondrial membrane. p139–41 induced energy-dependent swelling of isolated mitochondria by increasing inner membrane permeability to small cations (Na+, K+) and released Ca2+ from Ca2+-preloaded mitochondria. These effects as well as the ability of full-length p13II to alter mitochondrial morphology in cells required the presence of four arginines, forming the charged face of the targeting signal. The mitochondrial effects of p139–41 were insensitive to cyclosporin A, suggesting that full-length p13II might alter mitochondrial permeability through a permeability transition pore-independent mechanism, thus distinguishing it from the mitochondrial proteins Vpr and X of human immunodeficiency virus type 1 and hepatitis B virus, respectively. human T-cell leukemia virus type 1 circular dichroism cytochrome oxidase cyclosporin A mitochondrial inner membrane potential dinitrophenol 4-morpholinepropanesulfonic acid mitochondrial-targeting sequence open reading frame synthetic peptide spanning residues 9–41 of p13II permeability transition pore phosphate-buffered saline HTLV-11 is a complex retrovirus that is associated with two distinct pathologies, a leukemia/lymphoma of mature CD4+ T-cells and a neurodegenerative disease (tropical spastic paraparesis/HTLV-associated myelopathy). Adult T-cell leukemia/lymphoma and tropical spastic paraparesis/HTLV-associated myelopathy arise in a minority (2–5%) of HTLV-1-infected subjects after a latency period of years to decades. Although HTLV-1 was the first retrovirus demonstrated to cause neoplasia in humans, a number of aspects of its life cycle and pathogenic properties are still poorly understood (for recent reviews of HTLV-1, see Refs. 1Johnson J.M. Harrod R. Franchini G. Int. J. Exp. Pathol. 2001; 82: 135-147Crossref PubMed Scopus (89) Google Scholar and 2Green P.L. Chen I.S.Y. Knipe D.M. Howley P.M. Griffin D.E. Lamb R.A. Martin M.A. Roizman B. Straus S.E. Fields Virology. 4th Ed. Lippincott-Raven Publishers, Philadelphia2001: 1941-1969Google Scholar). The viral transcriptional regulator Tax is known to play a critical role in HTLV-1-associated cell immortalization through its ability to deregulate the expression of a vast array of cellular genes and interfere with cell cycle checkpoints. Although it is well established that HTLV-1 is oncogenic in vitro, neoplastic transformation of primary human lymphocytes cannot be attained by transducing Tax alone, suggesting that other viral (and host) determinants are also required for the emergence of adult T-cell leukemia/lymphoma. Furthermore, the low prevalence and long latency of leukemia in HTLV-1-infected individuals suggest the existence of mechanisms holding the oncogenic potential of the virus at bay, resulting in efficient adaptation to the host. Even less is known regarding the pathogenesis of tropical spastic paraparesis/HTLV-associated myelopathy, although a cytotoxic T-lymphocyte-mediated autoimmune mechanism has been postulated. To gain further insight into the mechanisms governing HTLV-1 replication and pathogenesis, a number of studies are currently focusing on recently described "accessory" proteins encoded in ORFs x-I and x-II, which are located in a 3′ portion of the viral genome termed the X region (3Berneman Z.N. Gartenhaus R.B. Reitz J.M.S. Blattner W.A. Manns A. Hanchard B. Ikehara O. Gallo R.C. Klotman M.E. Proc. Natl. Acad. Sci. U. S. A. 1992; 89: 3005-3009Crossref PubMed Scopus (133) Google Scholar, 4Ciminale V. Pavlakis G.N. Derse D. Cunningham C.P. Felber B.K. J. Virol. 1992; 66: 1737-1745Crossref PubMed Google Scholar, 5Koralnik I.J. Gessain A. Klotman M.E., Lo Monico A. Berneman Z.N. Franchini G. Proc. Natl. Acad. Sci. U. S. A. 1992; 89: 8813-8817Crossref PubMed Scopus (164) Google Scholar). Although the accessory ORFs are dispensable for virus propagation and immortalization of T-cells in vitro (6Derse D. Mikovits J. Ruscetti F. Virology. 1997; 237: 123-128Crossref PubMed Scopus (100) Google Scholar, 7Robek M.D. Wong F.-H. Ratner L. J. Virol. 1998; 72: 4458-4462Crossref PubMed Google Scholar, 8Collins N.D. D'Souza C. Albrecht B. Robek M.D. Ratner L. Ding W. Green P.L. Lairmore M.D. J. Virol. 1999; 73: 9642-9649Crossref PubMed Google Scholar), they are essential for efficient viral propagation in animal models (9Collins N.D. Newbound G.C. Albrecht B. Beard J.L. Ratner L. Lairmore M.D. Blood. 1998; 91: 4701-4707Crossref PubMed Google Scholar, 10Bartoe J. Albrecht B. Collins N.D. Robek M.D. Ratner L. Green P.L. Lairmore M.D. J. Virol. 2000; 74: 1094-1100Crossref PubMed Scopus (111) Google Scholar, 11Albrecht B. Collins N.D. Burniston M.T. Nisbet J.W. Ratner L. Green P.L. Lairmore M.D. J. Virol. 2000; 74: 9828-9834Crossref PubMed Scopus (87) Google Scholar). In addition, cell-mediated immune responses against these gene products have been demonstrated in infected individuals (12Pique C. Ureta-Vidal A. Gessain A. Chancerel B. Gout O. Tamouza R. Agis F. Dokhelar M.-C. J. Exp. Med. 2000; 191: 567-572Crossref PubMed Scopus (97) Google Scholar), thus providing strong evidence that these proteins are expressed during the natural history of HTLV-1 infection. We demonstrated that one of the gene products of the x-II ORF, named p13II, is selectively targeted to mitochondria. Expression of p13II results in specific alterations in mitochondrial morphology and distribution from an extensively interconnected network of string-like structures to clusters of spheroid bodies, suggesting that p13II might interfere with mitochondrial functionin situ (13Ciminale V. Zotti L. D'Agostino D.M. Ferro T. Casareto L. Franchini G. Bernardi P. Chieco-Bianchi L. Oncogene. 1999; 18: 4505-4515Crossref PubMed Scopus (85) Google Scholar). This observation along with the findings that the p13II-coding sequence is conserved among HTLV-1 isolates, that its mRNA is expressed during HTLV-1 infection and disease (5Koralnik I.J. Gessain A. Klotman M.E., Lo Monico A. Berneman Z.N. Franchini G. Proc. Natl. Acad. Sci. U. S. A. 1992; 89: 8813-8817Crossref PubMed Scopus (164) Google Scholar, 14Cereseto A. Berneman Z. Koralnik I. Vaughn J Franchini G. Klotman M.E. Leukemia (Baltimore). 1997; 11: 866-870Crossref PubMed Scopus (40) Google Scholar), and that cytotoxic T-lymphocytes against the x-II ORF are detected in HTLV-1-infected individuals (12Pique C. Ureta-Vidal A. Gessain A. Chancerel B. Gout O. Tamouza R. Agis F. Dokhelar M.-C. J. Exp. Med. 2000; 191: 567-572Crossref PubMed Scopus (97) Google Scholar) indicate that the protein might exert a significant biological role in viral replication and/or pathogenesis. The present study was undertaken to unravel the function of p13II through (i) determination of its submitochondrial localization and sequences required to induce the alterations in mitochondrial morphology and (ii) an assessment of the biophysical and biological properties of a set of synthetic peptides spanning residues 9–41, which include the amphipathic MTS of the protein. Results demonstrated the functional importance of a cluster of MTS arginines and indicated that the protein accumulates in the inner mitochondrial membrane. On the basis of observations made with the synthetic peptides, we propose that p13II increases mitochondrial permeability to small cations (e.g.Ca2+, Na+, and K+), causing swelling of the organelle and depolarization. Peptides were synthesized by solid phase methods and 9-fluorenilmethoxycarbonyl chemistry (15Fields G.B. Noble R.L. Int. J. Pept. Protein Res. 1990; 3: 161-214Google Scholar) using an Applied Biosystems (Foster City, CA) model 431 A peptide synthesizer (improved in our laboratories as described in Marin et al. (16Marin O. Meggio F. Sarno S. Pinna L.A. Biochemistry. 1997; 36: 7192-7198Crossref PubMed Scopus (46) Google Scholar). After cleavage with trifluoroacetic acid, the peptides were purified to homogeneity by reverse phase-high performance liquid chromatography on a Waters (Milford, MA) prepNova-Pak HR C18 column with a linear gradient of 10–45% acetonitrile at a flow rate of 12 ml/min. Peptide molecular weights were confirmed by mass spectroscopy using a matrix-assisted laser desorption ionization time-of-flight spectrometer (Maldi-1; Kratos-Shimadzu, Manchester, England). CD spectra were recorded with a Jasco (Easton, MD) J-170 spectropolarimeter. The instrument was calibrated with d-(+)-10-camphorsulfonic acid. Far-UV CD spectra were recorded at 25 °C at a peptide concentration of 15 μm in buffers specified in the legend to Fig. 2 using 0.1-cm path length quartz cells. The mean residue ellipticity [Θ] (deg·cm2·dmol−1) was calculated from the formula [Θ]MRW = (Θobs/10)MRW/lc), where Θobs is the observed ellipticity at a given wavelength, MRW is the mean residue weight of the peptide(s), l is the cuvette path length in cm, and c is the peptide concentration in g/ml. Lipid vesicles were prepared from dioleoylphosphatidylglycerol in trichloromethane. After evaporating the organic solvent under an argon stream, the dried lipid was dissolved in Tris/HCl buffer to obtain a final lipid concentration of 1.7 mm and sonicated for about 15 min until the solution became transparent. The p13II ORF was obtained from HTLV-1 molecular clone pCS-HTLV-1 (17Derse D. Mikovits L. Polianova M. Felber B.K. Ruscetti F. J. Virol. 1995; 69: 1907-1912Crossref PubMed Google Scholar) and cloned into vector pSG5 (Stratagene, La Jolla, CA), resulting in plasmid pSGp13II. Mutations in the p13II ORF were introduced by site-directed mutagenesis (QuikChange kit, Stratagene) and verified by cycle sequencing (fmol kit, Promega, Madison, WI). PCR amplifications were carried out using Vent DNA polymerase (New England Biolabs, Beverly, MA) in a GeneAmp 9600 thermal cycler (PerkinElmer Life Sciences). Restriction enzymes were purchased from New England Biolabs and Roche Molecular Biochemicals, and synthetic oligonucleotides were purchased from Invitrogen. Plasmids were purified by chromatography (Jetstar kit, Genomed, Bad Oeynhausen, Germany) and transfected into HeLa cells or the HeLa-derived cell line Hltat (Ref.18Schwartz S. Felber B.K. Benko D. Mfenyo E.M. Pavlakis G.N. J. Virol. 1990; 64: 2519-2529Crossref PubMed Google Scholar; used due to their high transfection efficiency) by calcium phosphate coprecipitation. Crude preparations of mitochondria were obtained from Hltat cells as follows. One day after transfection with pSGp13II, cells were scraped into PBS, pelleted, and then resuspended in a mitochondrial isolation buffer consisting of 300 mm sucrose, 10 mm MOPS, pH 7.4, 1 mm EDTA, and 4 mmKH2PO4 (19Petit P.X. O'Connor J.E. Grunwald D. Brown S.C. Eur. J. Biochem. 1990; 194: 389-397Crossref PubMed Scopus (213) Google Scholar) with 200 μmphenylmethylsulfonyl fluoride and 10 μg/ml leupeptin added as protease inhibitors. The suspension was passed 10 times through a 26-gauge needle and centrifuged 2 times at 625 × g for 10 min in a swinging bucket rotor to remove intact cells and large debris. The clarified supernatant was centrifuged at 17,500 ×g for 15 min; the resulting pellets containing mitochondria were resuspended in mitochondrial isolation buffer and recentrifuged. For Triton X-114 fractionation assays, mitochondrial pellets were resuspended in 10 mm Tris, pH 7.5, 150 mm NaCl, 200 μm phenylmethylsulfonyl fluoride, 10 μg/ml leupeptin, and 1% Triton X-114 and processed as described by Bordier (20Bordier C. J. Biol. Chem. 1981; 256: 1604-1607Abstract Full Text PDF PubMed Google Scholar) with the exception that all volumes were reduced by one-half; supernatants and pellets were solubilized and analyzed by SDS-PAGE/immunoblotting. Sodium carbonate extraction (21Fujiki Y. Hubbard A.L. Fowler S. Lazarow P.B. J. Cell Biol. 1982; 93: 97-102Crossref PubMed Scopus (1381) Google Scholar) was performed as follows. Washed mitochondrial pellets were resuspended in 100 mm sodium carbonate, pH 11.5, and incubated on ice for 30 min. The alkaline mixture was then combined with enough 2.5m sucrose prepared in 100 mm sodium carbonate to yield a final sucrose concentration of 1.6 m. This suspension was placed in an ultracentrifuge tube and overlaid with 1.25m sucrose, 100 mm sodium carbonate followed by a layer of 250 mm sucrose, 100 mm sodium carbonate and centrifuged at 47,000 rpm for 2 h in an SW60.1 rotor (Beckman Instruments). The step gradient was collected in 2 fractions, the 0.25–1.25 m sucrose interface plus the 1.25m sucrose layer, containing membrane-integrated proteins, and the 1.6 m sucrose layer, containing proteins not inserted into membranes. The fractions were diluted with water to reduce the sucrose concentration to 500 mm, and proteins were precipitated with 12% trichloroacetic acid, solubilized, and separated by SDS-PAGE/immunoblotting. Digitonin fractionation assays were performed by resuspending mitochondria in a sucrose-based solution (250 mm sucrose, 10 mm HEPES, pH 7.1, 10 mm KCl, 1.5 mm MgCl2, 1 mm EDTA, 1 mm EGTA, 200 μmphenylmethylsulfonyl fluoride, and 10 μg/ml leupeptin) with or without 0.5% digitonin; after incubation for 1 min on ice the suspensions were centrifuged at 17,500 × g for 15 min, and resulting supernatants and pellets were solubilized and analyzed by SDS-PAGE/immunoblotting. Solubilized proteins were separated in SDS-polyacrylamide gels (15% total; 37.5:1 acrylamide:bisacrylamide) along with a prestained protein marker (Cell Signaling, Beverly, MA) and then transferred to nitrocellulose (Sartorius, Goettingen, Germany). Blots were cut just above the 25-kDa marker; the top half was incubated with a mixture of goat anti-Hsp 60 (Santa Cruz, Heidelberg, Germany), mouse anti-Cox I (Molecular Probes, Leiden, Netherlands), and mouse anti-Porin (Calbiochem), and the bottom half was incubated with a rabbit serum raised against the carboxyl terminus of the x-II ORF (rabbit anti-Tof; 4). After incubation with appropriate horseradish peroxidase-conjugated secondary antibodies (Santa Cruz or Amersham Biosciences), the blots were developed using chemiluminescence detection reagents (Supersignal kit, Pierce) and exposed to Hyperfilm (Amersham Biosciences). For conventional electron microscopy, p13II-expressing Hltat cells were washed three times in PBS, pH 7.4, and fixed with PBS, 2% glutaraldehyde for 1 h at 4 °C. Samples were postfixed in 1% osmium tetroxide in Veronal acetate buffer, pH 7.4, for 1 h at 25 °C, stained with 0.1% tannic acid in the same buffer for 30 min at 25 °C and with uranyl acetate (5 mg/ml) for 1 h at 25 °C, dehydrated in acetone, and embedded in Epon 812. Thin sections were examined unstained or poststained with uranyl acetate and lead hydroxide. For immunoelectron microscopy, cells were fixed with a mixture of 4% paraformaldehyde and 0.2% glutaraldehyde prepared in 100 mm phosphate buffer for 2 h at 25 °C, washed, placed in 100 mmphosphate buffer containing 12% gelatin (Sigma), and solidified on ice. Gelatin blocks were infused with 2.3 m sucrose overnight at 4 °C, frozen in liquid nitrogen, and cryo-sectioned. Ultrathin cryo-sections were collected with sucrose and methyl cellulose and incubated with rabbit anti-Tof antibody diluted 1:10 in PBS, 1% bovine serum albumin. After several washes in PBS, 0.1% bovine serum albumin, the sections were incubated with 10-nm diameter colloidal gold particles conjugated with goat-anti rabbit IgG (British BioCell International, Cardiff, UK) diluted 1:20 in PBS. Control experiments were performed by omission of the primary antibody from the labeling procedure. After incubation with the antibodies, the cryo-sections were stained with a solution of 2% methyl cellulose and 0.4% uranyl acetate and examined by electron microscopy. Liver mitochondria isolated from albino Wistar rats weighing about 300 g were prepared by standard centrifugation techniques as described previously (22Costantini P. Petronilli V. Colonna R. Bernardi P. Toxicology. 1995; 99: 77-88Crossref PubMed Scopus (128) Google Scholar). Measurements of mitochondrial volume, membrane potential, and Ca2+ transport were carried out with a PerkinElmer Life Sciences 650–40 spectrofluorimeter equipped with magnetic stirring and thermostatic control. Mitochondrial volume was measured as the change in 90 ° light scattering at 540 nm (23Hunter D.R. Haworth R.A. Arch. Biochem. Biophys. 1979; 195: 453-459Crossref PubMed Scopus (591) Google Scholar, 24Petronilli V. Cola C. Massari S. Colonna R. Bernardi P. J. Biol. Chem. 1993; 268: 21939-21945Abstract Full Text PDF PubMed Google Scholar). Membrane potential and changes in extramitochondrial [Ca2+] were assayed by measuring the change in fluorescence intensity of rhodamine 123 (25Emaus R.K. Grunwald R. Lemasters J.J. Biochim. Biophys. Acta. 1986; 850: 436-448Crossref PubMed Scopus (722) Google Scholar) and calcium Green-5N (26Fontaine E. Ichas F. Bernardi P. J. Biol. Chem. 1998; 273: 25734-25740Abstract Full Text Full Text PDF PubMed Scopus (220) Google Scholar), respectively. Both probes were excited at 503 nm (2 nm slit), whereas emission was analyzed at 525 nm for rhodamine 123 and at 535 nm for calcium Green-5N, with the slit set at 5 nm in both cases. Details of assay conditions are provided in the legends to Figs. 5 and 6.Figure 6Effects of p13II peptides on membrane potential and Ca2+ fluxes in isolated mitochondria.Panel A, the experimental conditions were as in Fig. 5, panel A, but the medium was supplemented with 0.1 μm rhodamine 123. One milligram of rat liver mitochondria (RLM) was added followed by 1 μm wild-type p139–41 peptide (trace a); in trace b, the mitochondria were treated with 5 μmp139–41Q or p139–41P peptide followed by 200 μm DNP. Values on the ordinate refer to changes in rhodamine 123 fluorescence. In panel B, assays were performed at 25 °C in a 2-ml final volume containing 250 mm sucrose, 10 mm Tris-MOPS, 1 mmPi-Tris, 2 μg/ml CsA, 5 mm succinate-Tris, 2 μm rotenone, and 1 μm calcium Green-5N. The following additions (indicated by arrows) were made: 1 mg rat liver mitochondria (RLM), 10 μmCa2+, 0.2 μm ruthenium red (RR), 1 μm wild-type p139–41 peptide (trace a), 0.2 mm DNP (trace b) followed by 1 μm wild-type p139–41 peptide (last addition).View Large Image Figure ViewerDownload Hi-res image Download (PPT) 24–36 h after transfection, cells were washed twice with Dulbecco's modified Eagle's medium (DMEM) that had been prewarmed to 37 °C, fixed for 20 min with 3.7% formaldehyde in DMEM, permeabilized for 10 min with 0.1% Nonidet P40/PBS, and then incubated with the antibodies specified in the legend to Fig. 7. Signals were analyzed by confocal microscopy using a LSM510 microscope (Carl Zeiss, Jena, Germany) with argon (488 nm) and helium-neon (543 nm) laser sources and a 63× oil immersion objective; 0.9-μm optical slices were examined. Fig. 1A shows the 87-amino acid sequence of p13II, with its 10-amino acid MTS underlined. The PHDSec structure prediction program (27Rost B. Sander C. Proteins. 1994; 9: 55-72Crossref Scopus (1337) Google Scholar) suggests that the amino-terminal portion of the protein contains a short hydrophobic leader (amino acids 1–5) followed by an α-helix (amino acids 21–30) that includes the MTS (13Ciminale V. Zotti L. D'Agostino D.M. Ferro T. Casareto L. Franchini G. Bernardi P. Chieco-Bianchi L. Oncogene. 1999; 18: 4505-4515Crossref PubMed Scopus (85) Google Scholar). The latter region includes 4 arginine residues at positions 22, 25, 29, and 30, which are predicted to form a positively charged patch within the putative α-helix, thereby imparting amphipathic properties to this region (Fig.1B; Ref. 13Ciminale V. Zotti L. D'Agostino D.M. Ferro T. Casareto L. Franchini G. Bernardi P. Chieco-Bianchi L. Oncogene. 1999; 18: 4505-4515Crossref PubMed Scopus (85) Google Scholar). To directly verify these predictions, we generated synthetic peptides that included the p13II MTS and examined their structure by CD. Results showed that p139–41 (a peptide spanning amino acids 9–41 of wild-type p13II) failed to fold into an ordered structure when placed in an aqueous solution; in particular, the ellipticity minima at 208 and 222 nm that are characteristic of an α helix were not detected under these conditions (Fig.2A). Although the CD spectrum of the peptide changed when recorded in the organic solvent trifluoroethanol, utilized to promote the formation of an α-helical structure, the resulting conformation was predominantly unstructured. In contrast, the spectrum was dominated by the characteristic minima at 208 and 222 nm in the presence of 10 mm SDS, a concentration exceeding the critical micelle concentration, i.e. 0.8 mm under the assay conditions employed. The peptide also adopted an α-helical conformation when SDS was replaced with phospholipid vesicles, indicating that the MTS region is able to fold into an α helix only when exposed to a membrane-like environment. Fig. 2B shows the influence of the SDS concentration on the CD spectrum of peptide p139–41. Formation of an α helix became evident at an SDS concentration of 0.8 mm, i.e. the critical micelle concentration; the ellipticity minima were most pronounced at 1 mm SDS. As a prelude to subsequent functional studies, we next investigated the folding properties of peptides spanning residues 9–41 in which the 4 arginine residues at positions 22, 25, 29, and 30 were substituted with either 4 glutamines, alanine-leucine-leucine-alanine, or 4 prolines, resulting in peptides p139–41Q, AL, and P, respectively. While the glutamine and alanine-leucine substitution mutants were predicted to retain α helical folding, with just its positively charged face replaced with an uncharged polar cluster (mutant Q) or hydrophobic residues (mutant AL), the proline substitutions were predicted to disrupt the formation of an α helix. As shown in Fig. 2C, the spectra of peptides p139–41Q and p139–41AL exhibited minima at 208 and 222 nm in the presence of 10 mm SDS, indicating the formation of an α helix. In contrast, the p139–41P peptide failed to adopt an α-helical conformation. The finding that peptide p139–41 folded into an α helix when placed in a membrane-like environment prompted us to determine whether p13II is membrane-associated in the context of mitochondria. For this purpose, mitochondria isolated from human cells expressing the full-length p13II protein were extracted with Triton-X-114. This detergent is soluble in aqueous solutions at low temperatures but separates into aqueous and detergent phases above 20°C (20Bordier C. J. Biol. Chem. 1981; 256: 1604-1607Abstract Full Text PDF PubMed Google Scholar). Mitochondrial proteins partitioned in the aqueous and detergent phases were subjected to SDS-PAGE/immunoblotting and the distribution of p13II was compared with that of the soluble mitochondrial matrix protein Hsp 60 and the membrane proteins Porin and CoxI, which are inserted in the outer and inner mitochondrial membranes, respectively. As shown in Fig.3A, Hsp 60 was detected almost exclusively in the aqueous fraction, while Porin, CoxI, and p13II were found mainly in the detergent fraction, suggesting that p13II is likely to be associated with membranes rather than present in a soluble form in the matrix or intermembrane space. To test whether p13II was membrane-inserted, we subjected p13II-containing mitochondria to alkaline extraction with sodium carbonate (21Fujiki Y. Hubbard A.L. Fowler S. Lazarow P.B. J. Cell Biol. 1982; 93: 97-102Crossref PubMed Scopus (1381) Google Scholar) followed by ultracentrifugation through a sucrose step gradient. Fig. 3B shows immunoblots of the resulting soluble and membrane-bound fractions carried out to detect p13II, along with Hsp 60, CoxI, and Porin. p13II was detected mainly in the membrane-bound fraction along with the majority of CoxI (not evident in this exposure) and Porin, whereas Hsp 60 was detected mainly in the soluble fraction. Having obtained evidence that the majority of p13II is membrane-associated, we next determined whether the protein accumulates in the outer or inner membrane on the basis of its sensitivity to extraction by digitonin under isotonic conditions. As shown in Fig.3C, this treatment resulted in partial extraction of the outer membrane protein Porin; in contrast, neither the inner membrane protein CoxI nor p13II was extracted, thus providing evidence that p13II resides in the inner mitochondrial membrane. The immunoblots shown in Fig. 3 also revealed the presence of a p13II-related band of about 25 kDa (denoted by anasterisk) of unknown significance (see "Discussion"). To directly visualize the submitochondrial distribution of p13II and to examine its effect on mitochondrial morphology in more detail, electron microscopy was carried out on p13II-transfected human cells (Fig.4). Immunogold labeling to detect p13II (Fig. 4, panels A–E) revealed accumulation of gold particles inside mitochondria, mainly associated with the inner membrane and cristae. This observation supported the results obtained with the biochemical fractionation assays and indicated that the protein accumulates in the inner membrane. Ultrastructural examination of p13II-transfected cells revealed cells displaying different levels of morphological changes in their mitochondria (Fig. 4, panels F and G). Although some cells contained apparently unaltered mitochondria, others contained mitochondria showing fragmentation of the cristae and swelling, varying from slight disruption (see mitochondrion labeledM1) to substantial or almost complete loss of internal structures (mitochondria labeled M2 and M, respectively). Interestingly, mitochondria exhibiting more prominent alterations often appeared in close proximity to endoplasmic reticulum cisternae (see arrows and mitochondria labeled Min panel G). Although we could not positively identify p13II-expressing cells in this analysis, the percentage of cells showing altered mitochondria was consistent with the percentage of p13II-expressing cells determined by indirect immunofluorescence carried out on an aliquot of the transfected cells (results not shown), thus suggesting an association between p13II expression and the observed ultrastructural changes. The appearance of the altered mitochondria visualized by indirect immunofluorescence (13Ciminale V. Zotti L. D'Agostino D.M. Ferro T. Casareto L. Franchini G. Bernardi P. Chieco-Bianchi L. Oncogene. 1999; 18: 4505-4515Crossref PubMed Scopus (85) Google Scholar) and electron microscopy (Fig. 4, panels F and G) was suggestive of swelling, a phenomenon that arises from altered permeability of mitochondria to osmotically active species (for review, see Ref. 28Bernardi P. Physiol. Rev. 1999; 79: 1127-1155Crossref PubMed Scopus (1335) Google Scholar). This observation prompted us to assay the effects of the synthetic p13II peptides described above on isolated mitochondria. In the experiments reported in Fig.5, panel A, rat liver mitochondria energized with succinate were incubated in isotonic KCl, and the 90 ° light scattering of the suspension at 540 nm was monitored. The addition of 1 μm wild-type p139–41 peptide (Fig. 5, panel A, trace a) caused a rapid decrease in light scattering, which is indicative of mitochondrial swelling. After reaching a minimum level of light scattering, mitochondria underwent a partial recovery, and the signal eventually stabilized to an intermediate level. These volume oscillations, which were somewhat variable in extent among different mitochondrial preparations, are typical of the mitochondrial response to cation uptake through a channel, the contraction phase being due to transie
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