doi:10.1152/ajpheart.00833.2011 302:H167-H179, 2012. First published 28 October 2011;Am J Physiol Heart Circ Physiol

O'Connell, Rogerio F. Ribeiro, Jr., William C. Stanley and Kenneth WalshKyriakos N. Papanicolaou, Gladys A. Ngoh, Erinne R. Dabkowski, Kelly A.deathto ROS-induced mitochondrial dysfunction and cellmitochondrial fragmentation and improves tolerance Cardiomyocyte deletion of mitofusin-1 leads to

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Cardiomyocyte deletion of mitofusin-1 leads to mitochondrial fragmentationand improves tolerance to ROS-induced mitochondrial dysfunction and cell death

Kyriakos N. Papanicolaou,1 Gladys A. Ngoh,1 Erinne R. Dabkowski,2 Kelly A. O’Connell,2

Rogerio F. Ribeiro Jr.,2 William C. Stanley,2 and Kenneth Walsh1

1Whitaker Cardiovascular Institute, Boston University School of Medicine, Boston, Massachusetts; and 2Division ofCardiology and Department of Medicine, University of Maryland, Baltimore, Maryland

Submitted 22 August 2011; accepted in final form 24 October 2011

Papanicolaou KN, Ngoh GA, Dabkowski ER, O’Connell KA,Ribeiro RF Jr, Stanley WC, Walsh K. Cardiomyocyte deletion ofmitofusin-1 leads to mitochondrial fragmentation and improves tol-erance to ROS-induced mitochondrial dysfunction and cell death. AmJ Physiol Heart Circ Physiol 302: H167–H179, 2012. First publishedOctober 28, 2011; doi:10.1152/ajpheart.00833.2011.—Molecularstudies examining the impact of mitochondrial morphology on themammalian heart have previously focused on dynamin related pro-tein-1 (Drp-1) and mitofusin-2 (Mfn-2), while the role of the othermitofusin isoform, Mfn-1, has remained largely unexplored. In thepresent study, we report the generation and initial characterization ofcardiomyocyte-specific Mfn-1 knockout (Mfn-1 KO) mice. Usingelectron microscopic analysis, we detect a greater prevalence of small,spherical mitochondria in Mfn-1 KO hearts, indicating that the ab-sence of Mfn-1 causes a profound shift in the mitochondrial fusion/fission balance. Nevertheless, Mfn-1 KO mice exhibit normal left-ventricular function, and isolated Mfn-1 KO heart mitochondria dis-play a normal respiratory repertoire. Mfn-1 KO myocytes areprotected from mitochondrial depolarization and exhibit improvedviability when challenged with reactive oxygen species (ROS) in theform of hydrogen peroxide (H2O2). Furthermore, in vitro studiesdetect a blunted response of KO mitochondria to undergo peroxide-induced mitochondrial permeability transition pore opening. Thesedata suggest that Mfn-1 deletion confers protection against ROS-induced mitochondrial dysfunction. Collectively, we suggest thatmitochondrial fragmentation in myocytes is not sufficient to induceheart dysfunction or trigger cardiomyocyte death. Additionally, ourdata suggest that endogenous levels of Mfn-1 can attenuate myocyteviability in the face of an imminent ROS overload, an effect that couldbe associated with the ability of Mfn-1 to remodel the outer mito-chondrial membrane.

mitochondrial dynamics; GTPase; dynamin; membrane permeability;apoptosis; cardiomyocytes; reactive oxygen species

MERGING AND PARTITIONING the mitochondrial membranes areongoing processes that govern the structure and morphology ofthe organelle in a wide range of organisms and cell types (8,48). Extensive mitochondrial fusion and matrix continuity arecurrently viewed as adaptive mechanisms that allow mitochon-dria to cope with increased energetic demands and cellularstressors (60, 67). Furthermore, coupling selective fusion withfission is believed to constitute a means for mitochondrialquality control (68). In mammals, fusion of the outer mito-chondrial membrane (OMM) between adjacent mitochondria iscontrolled by mitofusins (Mfn-1 and Mfn-2; Refs. 10, 36, 56).Disrupting mitochondrial fusion via Mfn-1 or Mfn-2 ablationalters the distribution and morphology of mitochondria and

precipitates a stochastic process of mitochondrial dysfunc-tion (9, 10). Recently, the requirement of mitochondrial fu-sion for normal neuronal function and for mitochondrial DNA(mtDNA) maintenance in skeletal muscle has been docu-mented in mice (11, 12).

Despite the widespread utilization of fusion by mitochon-dria, the mechanism of membrane merging (i.e., tethering,formation of a fusion pore and lipid rearrangement) and howmitofusins participate in these processes remain incompletelyunderstood. In this regard, it has been shown that the C-termi-nal coiled-coil region of mitofusins is necessary in the initialtethering between opposing mitochondria and that guanosinetriphosphate (GTP) hydrolysis during mitofusin engagement isessential for subsequent OMM fusion steps (32, 43). Morerecently, it has been shown that the two isoforms are able tocomplement each other in the fusion process such that theformation of Mfn-1:Mfn-2 heterologous complexes can behighly effective in promoting fusion in vitro (24). The effect ofmitofusins in membrane fusion is counterbalanced by dy-namin-related protein-1 (Drp-1), a mediator of mitochondrialdivision (61, 70). Although mitofusins and Drp-1 catalyzeopposite reactions, they are structurally similar in that theyencompass a GTPase domain, and the mutation of key residuesin these domains can significantly alter the dynamics of mito-chondrial morphology in cells (46, 56, 57, 62).

Excessive mitochondrial fragmentation, typified by the pres-ence of numerous small and spherical mitochondria, is an earlyevent occurring during regulated cell death (apoptosis) (14, 39)and has been described in experimental models of heart failure(55). Reversing mitochondrial diminution via mitofusin activationor Drp-1 inhibition can prevent the progression of cell death insome contexts (19, 28, 35, 49, 64) but not in others (6, 52, 58). Onthe other hand, fragmentation alone is probably not sufficient totrigger cell death, because many cells harboring extensively frag-mented mitochondrial networks maintain apparent viability (10,31, 36, 37). Thus the question of whether mitochondrial fragmen-tation is sufficient to affect apoptosis remains controversial (53,63). Furthermore, it has been suggested that mitochondria-shaping proteins can influence cell death progressionthrough activities that do not primarily involve mitochon-drial fragmentation (34). For example, the mitochondrialfission factor Fis-1 has been shown to relay apoptosis-inducing signals before fragmentation occurs (27).

In heart, mitochondria are abundantly present where theyform a crystalline pattern and display limited motility (69).However, cardiac mitochondria appear capable for interorgan-ellar communication through the coordination of membranepotential (3, 5). To better understand the role of mitochondrialdynamics in cardiac myocytes, we have undertaken a mouse

Address for reprint requests and other correspondence: K. Walsh, MolecularCardiology/Whitaker Cardiovascular Institute, Boston Univ. School of Medi-cine, 715 Albany St., W611, Boston, MA 02118 (e-mail: kxwalsh@bu.edu).

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genetic approach to ablate mitofusins in cardiac myocytes.Previously, we constructed and characterized mice lackingMfn-2 in cardiac myocytes (50). This modification led to heartsthat displayed modest cardiac hypertrophy and mild functionaldeterioration. Mfn-2-deficient mitochondria were pleiomorphicand characteristically enlarged, and myocytes from these heartswere protected from mitochondrial permeability transition pore(MPTP) opening and cell death. Here, we constructed micelacking Mfn-1 in cardiac myocytes. These hearts are structur-ally and functionally normal. In marked contrast to Mfn-2-deficient hearts, however, mitochondria in Mfn-1-deficienthearts are smaller, yet myocytes are protected from reactiveoxygen species (ROS)-induced death. Furthermore, the assess-ment of isolated mitochondria showed greater resistance totert-butyl hydroxyperoxide (tBH)-induced MPTP opening.These findings highlight a striking functional difference be-tween Mfn-1 and Mfn-2 in controlling mitochondria size inmyocytes but also reveal considerable parallels regarding theinvolvement of mitofusins in regulating the mitochondrialresponse against ROS-induced dysfunction. Finally, our datachallenge the notion that smaller mitochondria predispose tostress-induced cell death.

MATERIALS AND METHODS

Animals. Mice carrying the Mfn-1loxP allele (stock Mfn1tm2Dcc;identification no. 029901-UCD) were obtained from the MutantMouse Regional Resource Center at University of California, Davis,and their generation has been previously described (11). These micewere mated with Myh6-cre transgenic mice designed to express crerecombinase selectively in cardiac myocytes and to drive site-specificrecombination of loxP sequences as early as embryonic day 9.5 (1,21). Mice with the genotype Mfn-1flox/flox;Myh6-cre�/� are termedMfn-1 KO. Mice with the genotype Mfn-1flox/flox;Myh6-cre�/� (lit-termates) or Mfn-1�/�;Myh6-cre�/� are collectively termed Mfn-1wild-type (WT) and were used interchangeably in the present study.The genetic background of the mice is 129S/C56BL6/BlackSwiss.The mice were housed in a 12-h light-dark cycle and temperature-controlled room and all handling procedures conformed to the regu-lations of and were approved by the Institutional Animal Care and UseCommittee of Boston University School of Medicine.

Echocardiography. The procedure to obtain echocardiograms andmeasurements from lightly anesthetized mice using the Vevo770system (http://www.visualsonics.com/vevo770) has been describedpreviously (50). Cardiac mass (CM) was calculated from long axisviews of the myocardium (epicardium and endocardium) in diastole(d) and systole (s) according to the equation:

CM � 1.05 � ��5

6Epicardialarea:d � EpicardialMajor:d � T�

� �5

6Endocardial area: d � Endocardial Major: d��

where T is the average wall thickness and 1.05 is the specific gravityconstant of the myocardium. The volumes in end-diastole and end-systole (EDV and ESV, respectively) were calculated according toSimpson’s equation that utilizes the area of the chamber at four levelsperpendicular to the long axis. These volumes were used to calculatethe ejection fraction according to the equation

EF � 100 ��EDV � ESV�

EDV

The fractional shortening (FS) was calculated from M-mode views ofthe myocardium at the papillary muscle level (short axis view)according to the equation:

FS � 100 ��LVID: d � LVID: s�

LVID: d

where LVID is the LV internal diameter.Electron microscopy. Rod-shaped pieces (�1 mm in diameter and

2 mm in length) were obtained from the LV free wall of adult malemice, fixed in Karnovsky’s solution (2% paraformaldehyde, 2.5%glutaraldehyde, and 0.1 M phosphate buffer; Electron MicroscopySciences) and processed until embedding into epon plastic (Embed-812; Electron Microscopy Sciences), as previously described (50).Ultra-thin sections containing predominantly longitudinal myofiberswere stained with 1% (wt/vol) uranyl acetate for 20 min followed byReynold’s lead citrate for 5 min. Micrographs were collected using aPhilips CM12 transmission electron microscope at 120 kV and�6,300 magnification. Scanned micrographs were analyzed usingImageJ software to manually generate masks of mitochondrial con-tours that were used for the calculation of mitochondrial area, maxi-mum mitochondrial diameter, mitochondrial perimeter, and total mi-tochondrial number.

Isolation of adult cardiac myocytes. Adult cardiac myocytes(ACMs) were isolated from hearts of male mice weighting 30 g ormore, as previously described (50, 51). Briefly, the hearts wereexcised and perfused through the aorta with a buffer containing thefollowing (in mM): 115 NaCl, 4.7 KCl, 0.6 KH2PO4, 0.6 Na2HPO4,1.2 MgSO4-7H2O, 0.032 phenol red, 10 KHCO3, 10 HEPES, 12NaHCO3, 20 glucose, 30 taurine, and 10 butanedione monoxime, pH7.46, at 37°C. For digestion of the extracellular matrix, the perfusionbuffer was supplemented with 19.5 units of Liberase TH (Roche), 0.5pM EDTA-free trypsin (Invitrogen), 25 �M CaCl2, and 25 �Mblebbistatin (Sigma). Ventricles were gently dissociated into rod-shaped myocytes that were suspended into perfusion buffer containingFBS (Hyclone) and were gradually reintroduced to CaCl2. For theirplating, myocytes were suspended in culture medium containingMEM (Invitrogen; contains 1.8 mM CaCl2) supplemented with 1%insulin-transferrin-selenium, 2 mM L-glutamine, 4 mM NaHCO3, 10mM HEPES, 2% penicillin/streptomycin, 5% FBS, 0.2% bovine calfserum, and 25 �M blebbistatin and seeded onto dishes previouslycoated with mouse laminin (BD Biosciences). After an initial platingstep for 1 h at 37°C and 2% CO2, the nonadherent cells were aspiratedand fresh culture medium was added for an additional 2-h periodbefore the beginning of treatments.

Confocal microscopy and image processing. Single myocytes wereseeded on glass bottom dishes (MatTek) that were previously coatedwith laminin and incubated in culture medium supplemented with 100nM tetramethylrhodamine methylester (TMRM; Molecular Probes)for 15 min at 37°C. That medium was replaced with TMRM-freemedium, and myocytes were imaged with a Zeiss LSM 710 confocalmicroscope using the Plan apochromat �63, 1.40 Oil DIC objective.A myocyte area of 67.48 � 67.48 �m was blindly selected for eachdish for continuous imaging. Illumination of TMRM was performedwith the 543 nm laser (the pinhole set at 50 �m), and fluorescence wasdetected between 554 and 652 nm. The laser power varied between0.2 to 12% for Mfn-1 WT (means: 2.7 � 0.8) and between 0.2 to 14%for Mfn-1 KO (means: 2.8 � 1.0). Following initial imaging, H2O2

was injected into the dish at a final concentration of 200 �M andtime-lapse imaging commenced immediately at intervals of 2.4 s for30 min to monitor the change in fluorescence intensity. Time-lapseimages were analyzed offline using ImageJ software to generatemitochondrial depolarization curves within a region of 25 � 25 �mfor each myocyte assessed. The area under the curve was chosen as aquantitative measurement of mitochondrial depolarization.

Myocyte viability assay. Freshly isolated ACMs were prepared asdescribed above reintroduced to calcium and resuspended in MEM-based medium that contained 1.2 mM CaCl2, 12 mM NaHCO3, 1%penicillin/streptomycin, 2.5% FBS, and 25 �M blebbistatin. ACMswere seeded on laminin-coated dishes (12 �g laminin/35-mm dish) ata density of 375,000 rod-shaped myocytes per dish. After an initial

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plating step at 37°C and 2% CO2 for 1 h, nonadherent cells wereremoved and fresh medium containing 20 �M H2O2 or PBS wereadded to the remaining cells. Two hours later, cells were switched tomedium containing 0.04% trypan-blue solution (Sigma) for 10 minand multiple fields per dish were photographed using a camera-equipped light microscope. Counting of blue vs. nonblue myocyteswas performed with ImageJ to calculate a death score (percentage ofcells that stained blue) per field. For comparisons, distribution curveswere generated using multiple fields per genotype per treatment, oralternatively the average death score per animal per genotype pertreatment was used.

Mitochondrial isolation and assessment of MPTP. Subsarcolemmalmitochondria (SSM) and interfibrillar mitochondria (IFM) were iso-lated from adult mouse hearts as previously described in detail (50).Size and membrane potential of SSM and IFM was determined byflow cytometry as previously reported (13, 50). Calcium uptake andtBH-induced calcium release were assessed in isolated SSM and IFM.In short, mitochondria were resuspended in 2.0 ml calcium-free buffercontaining 100 mM KCl, 50 mM MOPS, 5 mM KH2PO4, 5 �MEGTA, 1 mM MgCl2, 5 mM glutamate, and 5 mM malate and assayedfor Ca2� uptake in a fluorescence spectrophotometer at 37°C. Tert-butyl hydrogen peroxide (400 mM) was infused at a rate of 0.2�l/min, and the concentration of free Ca2� in the medium wascalculated by monitoring the fluorescence of the Ca2� indicatorcalcium green-5N (CaGN-5N; Molecular Probes) with an excitationand emission of 488 and 530.

Measurement of mitochondrial enzyme activities. Activities ofcitrate synthase, isocitrate dehydrogenase, and medium-chain acylcoenzyme A (acyl-CoA) dehydrogenase were measured from hearthomogenates and isolated mitochondria as previously described (47).

Determination of total glutathione levels in heart. To determineglutathione levels in heart, a commercially available kit was used(Abcam) and the procedure was performed according to manufac-turer’s specifications. Briefly, 40 mg of heart tissue were rapidlycollected and homogenized in 200 �l in ice-cold assay buffer. Fol-lowing deproteination, the samples were neutralized and the superna-tant was assessed for total glutathione (reduced and oxidized). Quan-tification of glutathione in the samples was performed spectrophoto-metrically, and the standard curve that was generated ranged from 4ng to 6 �g of glutathione.

Western blotting and quantitative real-time PCR. The proceduresfor the isolation, separation, transfer to membranes, and detection of

proteins (20 �g) from whole cardiac extracts were previously de-scribed (50). The antibodies used in the present study (diluted at1,000:1 in 3% blocking solution) include the anti-Mfn-1 (Universityof California, Davis/NIH; NeuroMabs; clone no. N111/24), anti-Mfn-2 (Sigma, N-terminal-specific), anti-Drp-1 (BD TransductionLaboratories), anti-cre (Novagen), anti-GAPDH (Cell Signaling),anti-porin/VDAC1 (Abcam), anti-uncoupling protein-3 (Thermo Sci-entific), and anti-FOF1-ATPase subunit-� (Invitrogen). For quantita-tive real-time PCR, total RNA was extracted from mouse hearts withfibrous-tissue mini kit (Qiagen) and 840 ng RNA were reversedtranscribed into cDNA using Quantitect reverse transcription kit(Qiagen). Quantification of cDNAs was performed with the SYBRgreen reagent (Applied Biosystems) and the StepOne real time sys-tem. Primers were designed using Primer3 and evaluated for genespecificity using BLAST. GAPDH was used as the housekeepinggene.

Statistical analysis. Handling and representation of the data wereperformed using SPSS software. All values shown are means � SE.For comparisons between two groups (i.e., WT vs. KO), the indepen-dent-samples two-tailed t-test was used (nonpaired). For comparisonsbetween four groups (i.e., two genotypes and two treatments), one-way ANOVA was used followed by the least significant differencespost hoc test. Differences were considered statistically significant forP � 0.05 or below. To compare the distribution curves betweengenotypes, the Kolmogorov-Smirnov test was applied.

RESULTS

Mfn-1 ablation and transcription pattern analysis. Micewith targeted deletion of Mfn-1 in cardiac myocytes are viableand survive through adulthood, in agreement with earlierobservations with globally deleted Mfn-1 mutant mice (11).Western blot analyses showed a complete absence of Mfn-1from cardiac extracts of Mfn-1 KO mice, suggesting an effec-tive abrogation of the Mfn-1 gene (Fig. 1, A and B). This eventdid not lead to appreciable changes in the level of Drp-1protein (Fig. 1A) or Mfn-2 (Fig. 1B). Despite the deletion ofMfn-1, the hearts did not display significant gross abnormali-ties (Fig. 1C). Furthermore, there was no reproducibly detect-able hypertrophy when normalizing the total heart weight totibia length (Fig. 1D; P 0.291).

Fig. 1. Characteristics of the mitofusin-1(Mfn-1) knockout (KO) heart. A: Westernblot analysis performed to detect endoge-nous levels of Mfn-1 and dynamin-relatedprotein-1 (Drp-1) in mouse heart extracts.GAPDH is used as a loading control. B: West-ern blot analysis performed to detect cre-specific reduction of Mfn-1 in the heart.Porin/VDAC1 is used as a mitochondrialoading control (WT, wild-type; HET,heterozygous for Mfn-1flox; KO, homozy-gous for Mfn-1flox and positive for cre).C: gross appearance of hearts from 5-mo-oldMfn-1 WT and KO mice; scale bar 2 mm.D: size of hearts is reported as the ratio ofheart weight (HW) to tibia length (TL; n 6WT; 4 mice were homozygous Mfn-1flox

without cre and 2 were homozygous Mfn-1�/� with cre and n 5 mice homozygousMfn-1flox with cre).

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To identify cellular pathways likely to be affected by thedeletion of Mfn-1 in hearts, quantitative real-time PCR analy-sis was performed and the results are reported in Table 1. Asshown, Mfn-1-deficient hearts displayed a 95% reduction inMfn-1 mRNA, while the mRNAs of other classical (Mfn-2,Drp-1, and Opa-1) and recently described (Mtp-18 and Slp-2)mitochondrial fusion/fission mediators decreased slightly orremained unchanged. Transcripts encoding mitochondria bio-genesis regulators (i.e., Tfam and Pgc-1�) did not changesignificantly, while genes encoding for electron transport chain(ETC) components such as Ndufb-5, Nd-5, and Cox4-1 weresignificantly attenuated. However, this was not a general fea-ture because the transcription of other ETC and ATPase sub-units was not significantly affected (i.e., Cox4–2, Cox-5b, andAtp-5o; Table 1). Finally, the mRNAs of Anp and Bnp weremarkedly elevated, but those of -Mhc and �-skeletal muscleactin were unchanged.

Mfn-1-deficient adult myocytes contain small and sphericalmitochondria. Analysis of the ultramicroscopic structure ofhearts from adult Mfn-1 WT and Mfn-1 KO mice detected the

formation of small ovoid or spherical mitochondria whenMfn-1 was genetically removed (compare Fig. 2A with Fig. 2Bor Fig. 2C with Fig. 2D). Although the presence of fragmentedmitochondria was quite prominent in most fields, in some areasof Mfn-1 KO myocytes the mitochondria assumed differentmorphologies to form clusters of lamellar often interdigitatingmitochondria with poorly defined boundaries (Fig. 2E). Thispattern was not observed in any Mfn-1 WT sections. Detailedanalysis of the mitochondrial dimensions and averaging over alarge population of mitochondria demonstrated a significantreduction in the cross sectional area of individual mitochondriaper field examined in Mfn-1 KO samples (Fig. 2F; P � 0.001).Despite these unusual structural features, the overall mitochon-drial mass estimated by mitochondrial volume density analysiswas not different between Mfn-1 WT and KO samples (Fig.2G, mitochondria; P 0.176). With the use of the sameanalysis, a small but significant decrease in the myofibrilvolume density was detected (Fig. 2G, myofibrils; P � 0.01).

Frequency distribution curves, constructed for maximummitochondrial diameter, were shifted to the left when Mfn-1was ablated, consistent with the recurrent presence of smallermitochondria in Mfn-1 KO hearts (compare Fig. 2H with Fig.2I for Mfn-1 WT and KO tissue, respectively; P � 0.001).Similar observations could also be made regarding the mito-chondrial perimeter (compare Fig. 2J with Fig. 2K, WT andKO tissue, respectively; P � 0.001). Another feature thatemerged from this analysis was the marked increase in thenumber of identifiable mitochondria when Mfn-1 was deleted(estimated number: 58 vs. 98 mitochondria/100 �m2 in WTand KO hearts, respectively). The detection of smaller mito-chondria that occur in higher numbers is in agreement with themeasurements showing that the overall mitochondrial volumedensity does not differ between WT and Mfn-1 KO samples.

Function of Mfn-1 KO hearts closely resembles that ofMfn-1 WT. The functional significance of Mfn-1 ablation incardiac myocytes was assessed in adult mice using noninvasiveechocardiography. The experimental groups included 11Mfn-1 WT and 13 Mfn-1 KO male mice that were �18 wk oldat the time of echocardiography (Table 2). The heart rateduring echocardiography was not significantly different be-tween Mfn-1 WT and KO mice (results not shown). Thecardiac mass calculated from B-mode long axis views of theLV was similar in hearts with or without Mfn-1, and this wasalso the case after correcting for body weight. Diastolic andsystolic LV volumes were not affected by Mfn-1 deletion, andas a result, the ejection fraction and the cardiac output wereindistinguishable between Mfn-1 WT and KO mice (Table 2).The fractional shortening, calculated from M-mode tracings,was also similar between groups. The peak aortic and pulmonicvalve flows did not differ significantly in the Mfn-1 KO hearts.Finally, examination of diastolic function in Mfn-1 WT andKO hearts did not reveal any differences (e.g., early mitralinflow, E; Table 2), suggesting that the overall systolic anddiastolic heart function is unlikely to be affected by theablation of Mfn-1 from myocytes.

Rate of mitochondrial depolarization downstream of ROSchallenge is decreased in Mfn-1 KO myocytes. We havepreviously demonstrated that the ablation of Mfn-2 in cardiacmyocytes is associated with a marked delay in the loss ofmitochondrial membrane potential (��m) (50). To examinethis

Table 1. Transcriptional regulation of relevant genes

TranscriptFold Induction/Reduction

(Mfn-1 KO vs. WT) P Functional Class

Mfn-1 0.074 0.001 Mitochondrial fusion/fissionMfn-2 0.838 0.017Opa-1 0.796 0.040Drp-1 0.805 0.041Mtp-18 1.047 0.587Slp-2 1.028 0.721Tfam 0.516 0.098 Mitochondrial biogenesisPgc-1� 0.409 0.078Err-� 1.250 0.307Endo-G 0.768 0.018 MTDNA maintenancePol- 1.002 0.986Ndufb-5 0.744 0.040 ETC/ATP synthaseNd-5 0.647 0.033Cox4-1 0.609 0.026Cox4-2 0.718 0.090Cox-5b 0.937 0.261Atp-5o 0.640 0.067Anp 5.699 0.001 Stress indicatorsBnp 5.427 0.001-Mhc 0.793 0.058�-Sk.actin 1.100 0.738Sod-2 0.676 0.007 ROS detoxificationBcl-2 1.345 0.001 Apoptosis/autophagyBeclin-1 0.865 0.353LC-3b 0.843 0.126

Values represent fold reduction/induction in the mitofusin-1 (Mfn-1) knock-out (KO) group (n 5) relative to wild type (WT; n 4). Comparisons forstatistical significance were performed with two-tailed Student’s t-test. Mfn-2,mitofusin 2; Opa-1, optic atrophy 1 long variant; Drp-1, dynamin-relatedprotein 1; Mtp-18, mitochondrial protein 18; Slp-2, stomatin like protein 2;Tfam, transcriptional factor A mitochondrial; Pgc-1�, PPAR- coactivator 1�;Err-�, estrogen receptor related factor-�; Endo-G, endonuclease- ; MTDNA,mitochondrial DNA; Pol- , polymerase- ; Ndufb-5, NADH-dehydrogenase(ubiquinone) subcomplex 1, polypeptide 5; Nd-5, NADH-dehydrogenasesubunit 5, mitochondrially encoded; Cox4-1, cytochrome c oxidase, subunit 4,isoform 1; Cox4-2, cytochrome c oxidase, subunit 4, isoform 2; Cox-5b,cytochrome c oxidase subunit 5b; Atp5o, F1FO-ATPase complex, oligomycinsensitivity-conferring subunit, polypeptide 5; Anp, atrial natriuretic peptide;Bnp, brain natriuretic peptide; -MHC, -myosin heavy chain; �-Sk actin;�-skeletal actin; Sod2, superoxide dismutase, mitochondrial; LC-3b; microtu-bule associated protein 1 light chain 3b; ETC, electron transport chain; ROS,reactive oxygen species.

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parameter in myocytes lacking Mfn-1, ACMs were isolatedfrom Mfn-1 WT and KO mice and challenged with H2O2-induced stress (200 �M for 30 min). H2O2 has been exten-sively used to induce mitochondrial ROS overload, membranedepolarization, and ultimately cell death in myocytes (2, 38,66). The ability of mitochondria to retain the mitochondrial dyeTMRM was quantified by repetitive confocal imaging over a30-min period after H2O2 injection. The treatment with H2O2

and a concomitant photochemical ROS production due toTMRM illumination (72) effectively lead to mitochondrialdepolarization within intact myocytes, as shown in Fig. 3A.Compared with WT, Mfn-1 KO myocytes contained mitochon-dria that were refractory to depolarization (Fig. 3A, comparethe fluorescence intensity in the different time points). Whenplotted against time, the depolarization curve of Mfn-1 KOmyocytes significantly shifted to the right, indicating a delay in

this process (Fig. 3B). This was not due to alterations in basalmembrane potential (��m) because the TMRM fluorescencewas similar between the two groups at t0 (Fig. 3C). The abilityof mitochondria to retain TMRM was quantified as the areaunder the curve. This analysis showed that the TMRM reten-tion values are significantly higher in Mfn-1 KO myocytes(Fig. 3D; 11 WT or 15 KO myocytes from 7 different exper-iments; P � 0.01). To examine whether the ablation of Mfn-1alters the scavenging capacity for H2O2, we examined theexpression of genes related to glutathione, a central intracellu-lar H2O2 detoxifier. As shown in Fig. 3E, the ablation of Mfn-1did not result to consistent increases in genes regulating glu-tathione availability. Although glutathione peroxidase-1 wassignificantly increased, glutathione synthase was decreased.Additionally, the expression of the mitochondria-targeted glu-tathione peroxidase-4 was unchanged (Fig. 3E). Furthermore,

Fig. 2. Electron microscopy images showing mitochondria and myofibrils from adult hearts. A: 5-mo-old Mfn-1 WT. B: littermate Mfn-1 KO, the scale bars inA and B 2 �m. C: interfibrillar mitochondria in Mfn-1 WT heart. D: interfibrillar mitochondria in Mfn-1 KO heart, scale bars in C and D 0.5 �m. E: distinctarea of Mfn-1 KO heart (myofibrils present but not shown) containing unusually elongated/lamellar mitochondria; scale bar 0.5 �m. F: area per individualmitochondrion measured from electron micrographs taken from 2 animals per genotype. Numbers in circles show the number of fields used per genotype (***P �0.001). G: mitochondrial and myofibril volume density estimated by the grid method expressed as a percentage (grid contains 9 � 10 dots; **P � 0.01).H: histogram showing the distribution of maximum mitochondrial diameter of all the mitochondria identified (n 3,927) in WT samples. I: histogram showingthe distribution of maximum mitochondrial diameter of all the mitochondria identified (n 6,494) in KO samples. D is value for most extreme differencesdetermined by Kolmogorov-Smirnov analysis (P � 0.001) that the 2 histograms belong to the same distribution. J–K: same as in H and I for Mfn-1 WT andMfn-1 KO, respectively, only that the histograms were created using mitochondrial perimeter.

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direct quantification of glutathione levels in heart extracts didnot reveal any differences between groups (Fig. 3F). Milduncoupling of the ETC is thought to confer a protective effectin mitochondria undergoing ROS challenge (42, 66). To testwhether alterations in uncoupling proteins occur during Mfn-1deletion, we examined the protein levels of uncoupling pro-tein-3 (UCP-3). As shown in Fig. 3G, the levels of UCP-3 wereunaffected by the deletion of Mfn-1. Collectively, it can beconcluded from this analysis that Mfn-1 facilitates mitochon-drial depolarization downstream of H2O2, but this does notinvolve alterations in glutathione availability or ETC uncou-pling.

Mfn-1 KO myocytes are protected from ROS-induced celldeath. Irreversible membrane depolarization is closely associ-ated with mitochondrial dysfunction and cell death. Therefore,the delayed mitochondrial depolarization observed in Mfn-1KO myocytes could have an impact on myocyte survival. Toexamine myocyte viability, ACMs from Mfn-1 WT and KOmice were exposed to PBS or to H2O2-induced stress (20 �Mfor 2 h) to induce mitochondrial dysfunction and cell death(23). Myocyte death following these treatments was assessedmicroscopically on the basis of trypan-blue staining (Fig. 4A).Multiple fields per genotype per treatment were scored accord-ing to the percentage of trypan-blue positive cells (0–100%dead myocytes). In PBS-control conditions, the majority offields quantified in Mfn-1 WT and KO myocytes tended tocluster to the left part of the x-axis (Fig. 4B). However, inH2O2-treated conditions, this distribution collapses and spreadsrightward because many fields have a higher percentage ofdead cells. Although this occurred with the Mfn-1 WT myo-cytes, Mfn-1 KO myocytes had improved tolerance to H2O2

seen as a significant decrease in the number of dead cells (Fig.4B, compare the distribution of the red bars in the panels in theright; WT and KO). In agreement with these observations, themean percentage of trypan-blue myocytes in H2O2-treatedmyocytes was significantly different when comparing averages

between animals (Fig. 4C; P � 0.05 WT vs. KO and P � 0.05PBS vs. H2O2). Recent evidence suggests that Mfn-1 is tar-geted for degradation by the proteasome in response to intra-cellular stress (22, 65). In agreement with these observations,we find that the levels of Mfn-1 protein were attenuated inMfn-1 WT myocytes exposed to H2O2 (Fig. 4D). Thus theremoval of Mfn-1 may be advantageous for the cell duringstress, providing an explanation why myocytes rendered defi-cient in Mfn-1 (Mfn-1 KO) are more tolerant to ROS-relatedinsults.

Characteristics of isolated mitochondria. The protein yieldsof subsarcolemmal mitochondria (SSM) and interfibrillar mi-tochondria (IFM) did not decrease significantly upon Mfn-1ablation (Table 3), although the Mfn-1 KO IFM appeared tohave slightly higher protein content than WT (11.6 � 0.8 vs.14.1 � 0.78 mg mitochondrial protein/g wet mass respectively;P 0.051; Table 3). Further, the activities in whole tissuehomogenates were similar for the mitochondrial oxidativeenzymes citrate synthase (3.84 � 0.25 vs. 4.88 � 0.48�mol·mg protein�1·min�1 for WT and KO, respectively),isocitrate dehydrogenase (0.60 � 0.15 and 0.85 � 0.14�mol·mg protein�1·min�1), and medium chain acyl-CoA de-hydrogenase (0.28 � 0.02 and 0.31 � 0.03 �mol·mgprotein�1·min�1), consistent with the notion that the overallmitochondrial mass did not change in Mfn-1 KO hearts (forexample, Fig. 2G). Mitochondrial size was found to be smallerby flow cytometry for both SSM and IFM in the Mfn-1 KOhearts than in WT (Table 3), consistent with the electronmicroscopy analysis (Fig. 2F). The membrane potential ofSSM and IFM was not affected by Mfn-1 deletion (Table 3).Mitochondrial respiration was unaffected by deletion of Mfn-1,with the exception of a greater state III respiration in the IFMfrom Mfn-1 KO with succinate � rotenone and a higherrespiratory control ratio with glutamate � malate (Table 3).Thus deletion of Mfn-1 resulted in smaller mitochondria butdid not have a deleterious effect on mitochondrial content oroxidative capacity.

ROS-induced MPTP opening is decreased in isolated inter-fibrillar mitochondria. The ability to uptake Ca2� is used toassess the physiological state of mitochondria (7). As shown inFig. 5A, Mfn-1 KO SSM exhibit a similar Ca2� uptake as WTSSM. On the other hand, the same assay detected a modestreduction in the ability of Mfn-1 KO IFM to uptake extra-mitochondrial Ca2� (Fig. 5B). To assess the ability of mito-chondria to undergo ROS-induced MPTP, we utilized thecontinuous infusion of tBH. Addition of tBH triggered therapid release of Ca2� from mitochondria, which was notdifferent between Mfn-1 WT and KO SSM for the duration ofthe assay (Fig. 5C) but was found to be significantly blunted inMfn-1 KO IFM (Fig. 5D), consistent with the resistance toROS-induced mitochondrial dysfunction and death observed inisolated myocytes (Figs. 3 and 4).

DISCUSSION

Here we report the generation and functional characteriza-tion of mice that lack the mitochondrial fusion protein Mfn-1in cardiac myocytes. Mice lacking Mfn-1 exhibited normalheart function, but the mitochondria within Mfn-1-deficientmyocytes were small and spherical. These findings are inmarked contrast to our previous analysis of Mfn-2-deficient

Table 2. Echocardiographic analysis

Mfn-1 WT (n 11) Mfn-1 KO (n 13)

Age, day 125 � 11 129 � 11BW, g 36.3 � 5.3 34.7 � 3.8Cardiac mass, mg 165.7 � 14.9 167.58 � 16.0CM/BW, mg/g 4.63 � 0.62 4.86 � 0.55EDV, �l 76.11 � 15.6 84.23 � 18.15ESV, �l 32.43 � 9.25 38.32 � 15.96EF, % 57.47 � 6.90 56.28 � 9.39CO, ml/min 22.53 � 4.70 22.79 � 3.59FS, % 56.65 � 9.19 55.23 � 11.80PAF, mm/s 1252 � 213 1403 � 315PPF, mm/s 842 � 136 820 � 123E, mm/s 583 � 126 513 � 101A, mm/s 447 � 95 420 � 86ET, ms 49.50 � 5.33 45.14 � 4.35E’, mm/s 11.89 � 2.41 11.36 � 2.29A’, mm/s 11.56 � 1.02 11.57 � 1.10

Results are means � SD. For derivation of the different parameters pleaserefer to MATERIALS AND METHODS. Comparisons for statistical significance wereperformed with two-tailed Student’s t-test. BW, body weight; CM, cardiacmass; EDV, end-diastolic volume; ESV, end-systolic volume; EF, ejectionfraction; CO, cardiac output; FS, fractional shortening; PAF, peak aortic flow;PPF, peak pulmonic flow; E, early mitral inflow; A, late mitral inflow; ET,ejection time; E’, early relaxation velocity at the mitral annulus; A’, laterelaxation velocity at the mitral annulus.

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Fig. 3. Mitochondrial depolarization downstream of H2O2

is delayed in single myocytes, but this does not involvealtered H2O2 scavenging or electron transport chain un-coupling. A: representative images of myocyte areas (�25�m) containing mitochondria undergoing depolarizationwith different kinetics (Mfn-1 WT above and Mfn-1 KObelow). H2O2 was injected at t0 and imaging beganimmediately. Images shown were taken at the indicatedtime points (minutes) following H2O2 injection. B: graphshowing dissipation of tetramethylrhodamine methylester(TMRM) fluorescence of a selected region of mitochon-dria after H2O2 exposure. On the y-axis, TMRM fluores-cence [arbitrary units (AU)] is a quantitative indicator ofinner membrane potential (��m) for the selected group ofmitochondria. On the x-axis, time after H2O2 injection isshown as consecutive slices (time interval: 2.4 s). Chan-nel 1 (blue) shows the course of depolarization in Mfn-1WT myocytes, and channel 2 (red) shows the depolariza-tion of Mfn-1 KO myocytes at the same conditions.C: TMRM fluorescence at t0 is quantified in myocytesfrom each group as a measure of basal membrane poten-tial (��m; P 0.189). D: capacity of mitochondria tolose TMRM is quantified as the area under the curve fromtraces such as those shown in B. Confocal microscopyexperiment was done using 4 WT and 3 KO mice thatwere 2.5–5 mo old. Number of myocytes assessed areshown in circles for each group (6 myocytes were Mfn-1flox without cre, 5 were Mfn-1� with cre, and 15 werehomozygous Mfn-1flox with cre). E: relative levels ofmRNAs encoding for proteins with the capacity to alterglutathione levels are quantified in whole-heart extracts.Gss, glutathione synthase; Gpx-1, glutathione peroxi-dase-1; Gpx-4, glutathione peroxidase-4, mitochondrialvariant. F: levels of total glutathione (reduced and oxi-dized) in whole heart extracts are shown in �g/mg for the2 experimental groups (2 mice were Mfn-1flox withoutcre, 2 mice were Mfn-1� with cre, and 4 mice werehomozygous Mfn-1flox with cre, aged 3.5–4 mo old).G: protein abundance of uncoupling protein-3 (UCP-3)and �-subunit of the FOF1-ATPase (FOF1-�) in Mfn-1WT and KO hearts. *P � 0.05, **P � 0.01.

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myocytes that displayed larger mitochondria (50). De-spite mitochondrial fragmentation, Mfn-1-deficient myocyteswere protected from ROS-induced mitochondrial depolariza-tion and cell death.

Why are mitochondria smaller in Mfn-1-deficient myo-cytes yet enlarged in myocytes that are deficient for Mfn-2?One possibility is that the degree of mitochondrial size isdependent on the relative levels of mitofusin GTPase activ-ity within the cell. Earlier studies (26) comparing mitofusinshave shown that the two homologs exhibit considerabledifferences in terms of binding and hydrolyzing GTP, whereMfn-1 has eightfold higher GTPase activity than Mfn-2.Furthermore, it has been shown that endogenous Mfn-1 caninteract with mutant Mfn-2 to form fusion-competent com-

plexes in cells, whereas endogenous Mfn-2 is inefficient inthis process (15). Collectively, these data suggest that Mfn-1is more effective in driving mitochondrial fusion thanMfn-2. Because it has been demonstrated that heterotypicinteractions between Mfn-1 and Mfn-2 are favored formitochondrial fusion compared with homotypic interactions(24), the balance of Mfn-1 relative to Mfn-2 may be animportant feature that determines mitochondrial size. In thisregard, it is possible that under certain conditions, thediminished GTPase activity of Mfn-2 may serve to retardthe fusion activity of Mfn-1 protein within the heterotypiccomplex. For example, if an Mfn-2 threshold is surpassed ina cell that expresses both isoforms, excessive Mfn-2 expres-sion could reverse the mitochondrial fusion activity of

Fig. 4. Viability assay of single adult cardiac myocytes. Experiment was done with 5 WT and 5 KO mice 2.5–5 mo old. A: microscopic fields containing singlemyocytes from WT and KO hearts previously treated in parallel with PBS or H2O2. Viable (unstained) vs. dead (blue) myocytes were counted, and each fieldwas scored for the percentage of blue cells (death score). B: distribution analysis of all the fields scored (shown by n) according to their death score (0–100%).Clustering at the left of the x-axis indicates a low death index. C: average death score per treatment and genotype (n 5 Mfn-1 WT or KO mice; *P � 0.05WT vs. KO and #P � 0.05 PBS vs. H2O2). D: Western blot analysis performed to detect Mfn-1 protein levels in Mfn-1 WT and KO single myocytes followingtreatment with PBS and H2O2. GAPDH is used as a protein loading control. Numbers at left indicate respective molecular masses (37–250 kDa).

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Mfn-1 by functioning in a manner that is analogous to adominant-negative isoform.

In the human and mouse heart, both Mfn-1 and Mfn-2 arehighly expressed (Ref. 56 and our unpublished observations),and it is conceivable that by deleting either of the mitofusinswill displace the ratio from its set point and increase/decreasethe efficiency of fusion. Consistent with this hypothesis, we(50) have previously generated mice with cardiomyocyte-specific deletion of Mfn-2. Lack of Mfn-2 resulted in formationof large globular mitochondria indicating that an excess ofMfn-1 activity (via removal of Mfn-2) leads to mitochondrialenlargement, possibly by enhancing fusion between mitochon-dria during biogenesis. Conversely, in the present study whereMfn-1 is ablated under identical conditions, we find thatmitochondria become smaller and spherical (Fig. 2), indicatingthat a relative excess of Mfn-2 activity can lead to mitochon-drial fragmentation. It is important to note that when onemitofusin is ablated, the levels of the other isoform do notchange significantly (Fig. 1B; Ref. 50). Our observation thatdeletion of mitofusins leads to distinct mitochondrial morphol-ogies is in concordance with earlier findings with mitofusin-deficient mouse embryonic fibroblasts (MEF; Ref. 10), whereit was noted that Mfn-1-deficient MEFs contain small sphericalmitochondria and Mfn-2-deficient MEFs contain large globularmitochondria. Similarly, the diminished GTPase activity ofMfn-2 may explain why Mfn-2 overexpression, in the presenceof endogenous Mfn-1, induces the breakdown of mitochondrialtubules into small spherical structures relative to the WTcondition (18, 25, 54, 57). In light of the above considerations,we suggest that in myocytes (and probably in other cells)excessive Mfn-1 activity leads to mitochondrial enlargement,whereas excessive Mfn-2 activity leads to mitochondrial dim-inution.

Notably, these shifts in Mfn-1/Mfn-2 balance and the ac-companying morphological transitions do not lead to signifi-cant impairment in the biochemical properties of mitochondria,and heart function is not severely impaired either in Mfn-1 KOor Mfn-2 KO mice. In this regard, we report here that Mfn-1KO hearts are functionally normal by echocardiography (Table

2) and cardiac hypertrophy is not detectable. However, in thecase of Mfn-2 KO mice the hearts were modestly hypertro-phied (50). Given the contrast in mitochondrial size betweenMfn-2 and Mfn-1 KO hearts, and the absence of hypertrophy inthe latter, it could be suggested that the modest hypertrophydetected in Mfn-2 KO hearts is related to the mitochondrialenlargement. More specifically, it might be that hypertrophy ofMfn-2 KO myocytes occurs as the consequence of an intracel-lular burden imposed to myocytes due to the inclusion ofaberrantly enlarged mitochondria. It has been previously re-ported (29) that experimental enlargement of mitochondriaalters the mechanical properties of cardiac myocytes. Further-more, the notion that mitochondria can cause cellular dysfunc-tion due to their increased size has been suggested for cere-bellar neurons (30). Interestingly, Mfn-2 conditionally defi-cient neurons are found to undergo degeneration in thecerebellum, and mitochondria in these cells form conglomer-ates within the soma that appear unable to enter the dendritesdue to their size (11). On the other hand, in that same study noabnormalities were detectable in Mfn-1-deficient mice, indi-cating that Mfn-1-deficient neurons are functionally intact. Inthis regard, it would be interesting to examine whether mito-chondria in Mfn-1 KO neurons have normal size or whetherthey are reduced in size as we detect in the Mfn-1 KO heart.

Mitochondrial dynamics appear to be intimately related tocell death pathways, but the details of this interaction remainlargely obscure. A common theme that emerges from the studyof cells undergoing apoptosis is that they exhibit mitochondrialfragmentation that occurs almost simultaneously with the re-lease of cytochrome c (41). Notably, it has been reported thatrepressing mitochondrial fragmentation by inhibiting Drp-1interferes but does not block the apoptotic process (17, 19, 28,52). Although it remains to be determined how smaller mito-chondria facilitate the apoptotic process, it is now clear thatmitochondrial fragmentation alone does not necessarily lead tocell death or any other apparent cellular dysfunction in severalcontexts (4, 59). Our current model provides an experimentalsystem to test the relationship between mitochondrial fragmen-tation and cell death induction in cardiac myocytes, since the

Table 3. Function of isolated cardiac mitochondria

Subsarcolemmal Mitochondria Interfibrillar Mitochondria

Mfn-1 WT (n 6) Mfn-1 KO (n 6) Mfn-1 WT (n 6) Mfn-1 KO (n 6)

Yield, mg mitochondrial protein/g wet mass, 18.2 � 1.0 16.5 � 5.8 11.6 � 0.8 14.1 � 0.78Mitochondria diameter, arbitrary units 534 � 13 490 � 6* 487 � 8 464 � 4*��m (JC-1 aggregate/monomer) 3.36 � 0.43 2.86 � 0.30 3.26 � 0.53 3.18 � 0.40Enzyme activities, nmol·mg mitochondrial protein�1 ·min�1

Citrate synthase 1.79 � 0.28 2.17 � 0.28 2.64 � 0.98 2.34 � 0.57Medium chain Acyl-CoA dehydrogenase 0.22 � 0.02 0.22 � 0.01 0.23 � 0.02 0.23 � 0.05Respiration, nmol·mg mitochondrial protein�1·min�1

Glutamate � malate: state III 97.2 � 6.6 95.4 � 4.4 99.5 � 3.8 117 � 7.3Glutamate � malate: state IV 39 � 3.1 33.4 � 2.3 46.7 � 2.7 39.3 � 2.3Glutamate � malate: RCR 2.55 � 0.25 2.94 � 0.27 2.15 � 0.11 3.00 � 0.20*Palmitylcarnitine � malate: state III 59.1 � 5.8 83.7 � 10 77.3 � 20 69.3 � 7.7Palmitylcarnitine � malate: state IV 24.9 � 0.93 31.5 � 2.7 32 � 5 28 � 2.8Palmitylcarnitine � malate: RCR 2.41 � 0.3 2.42 � 0.4 2.29 � 0.34 2.57 � 0.34Succinate � rotenone: state III 417.4 � 29.4 454.4 � 17.7 454.2 � 29.3 530.7 � 17*Succinate � rotenone: state IV 167.8 � 10.5 178.9 � 12.2 198.8 � 22.6 215 � 11.4Succinate � rtenone: RCR 2.37 � 0.07 2.66 � 0.1 2.20 � 0.05 2.56 � 0.15

Values are means � SE. Each data point (total of 6 per genotype) represents mitochondria pooled from 2 hearts. ��m, Basal membrane potential; JC-1,5,5=,6,6=-tetrachloro-1,1=,3,3= -tetraethylbenzimidazolylcarbocyanine iodide; RCR, respiratory control ratio *P � 0.05, between genotypes (Mfn-1 WT and KO)by unpaired two-tailed t-test within the same mitochondrial group (subsarcolemmal mitochondria and interfibrillar mitochondria).

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Mfn-1 KO myocytes acquire increased numbers of fragmentedmitochondria (Fig. 2). Accordingly, by subjecting these myo-cytes to ROS-induced stress we do not detect any evidence ofsensitivity towards cell death. In contrast, Mfn-1 KO myocytesare found to be protected against ROS-induced membranedepolarization (Fig. 3), and cell death is significantly attenu-ated in Mfn-1 KO myocytes exposed to H2O2 (Fig. 4). Takentogether, our findings are in agreement with the notion thatpreexisting mitochondrial fragmentation is not sufficient topredispose myocytes to death. More importantly, the ablationof Mfn-1 confers resistance against ROS-induced mitochon-drial depolarization and correlates well with a decreased rate ofcardiomyocyte death.

The acquired ability of Mfn-1 KO mitochondria to toler-ate deleterious amounts of H2O2 does not appear to be dueto a preexisting secondary activation of redox defense path-ways. In support of the above, we find that the levels ofglutathione and UCP-3 in heart do not differ significantly

between the two groups (Fig. 3, F and G). In addition, usingisolated mitochondria, we were able to detect resistanceagainst tBH-induced MPTP opening (Fig. 5), consistentwith a delayed mitochondrial depolarization observed withsingle myocytes (Fig. 3D). The experiments with isolatedmitochondria also revealed that the increased toleranceagainst ROS is likely to be more dominant or exclusivelylocalized to IFM rather than the SSM (Fig. 5, C and D), andadditional work will be required to delineate why thisproperty is selectively detected in this Mfn-1 KO mitochon-drial population. Nevertheless, the work with isolated mi-tochondria indicates that the ability of Mfn-1 KO myocytesto tolerate a ROS challenge is at least in part attributable toa blunted response of Mfn-1 KO mitochondria to undergoMPTP opening downstream of ROS overload.

Fusion between biological membranes, including those en-veloping mitochondria, has been previously thought of as aleakage-free process where the integrity of the rearranging

Fig. 5. Calcium-uptake capacity and reactive oxygen species (ROS) sensitivity of mitochondria isolated from Mfn-1 WT and KO hearts. A: rate of Ca2� uptakeby subsarcolemmal mitochondria (SSM) is monitored as a decrease in fluorescence of the extra-mitochondrial Ca2� indicator (CaGN-5N; 200 nM). B: decreasein CaGN-5N fluorescence over time due to the uptake of Ca2� by interfibrillar mitochondria (IFM). C: SSM mitochondria were primed for mitochondrialpermeability transition pore (MPTP) opening with a low concentration of Ca2� (22.5 �M) and were subsequently exposed to increasingly higher doses of tBH.Mitochondrial Ca2� release as a result of tBH-induced MPTP was monitored as an increase in CaGN-5N fluorescence. D: increase in CaGN-5N fluorescencedue to tBH-induced MPTP opening in IFM. AU, arbitrary fluorescence units; tBH, tert-butyl-hydroxyperoxide. *P � 0.05, between the 2 genotypes for theindicated time points.

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membranes is not breached (71). Nevertheless, there is exper-imental evidence to suggest that fusion may involve the for-mation of holes on membranes (16). For example, it has beenobserved that during the early steps of hemagglutinin-inducedfusion the membranes exhibit a transient increase in permea-bility that is resolved later as fusion progresses (20). Thisobservation is in agreement with theoretical models that predictthe opening of a hole on the area near the hemifusion stalk,(i.e., an intermediate structure that connects the apposed mem-branes during early fusion; Ref. 45). Breaching membraneintegrity is also predicted from calculations of the tensionforces that develop during the formation of the hemifusionstalk (33). Therefore, although membrane fusion is beneficialfor a number of cell processes that require content mixingbetween distinct compartments, it also entails a transient lossof membrane integrity to some extent. Along these lines, wehave previously reported that Mfn-2 can significantly contrib-ute to MPTP formation in cardiac myocytes treated with ROSand in the present study we extend these observations toMfn-1. Taken together, our observations suggest a previouslyunsuspected connection between mitochondrial fusion andstress-induced MPTP formation.

Apart from MPTP formation, which requires simultaneousopening of a channel-like pore in the IMM and a pore (or hole)on the OMM, permeabilization of the OMM alone (via forma-tion of lipidic pores) is also sufficient to trigger cell death.Along these lines, it has been recently reported that anothermediator of mitochondrial dynamics, Drp-1, can promote for-mation of holes on liposome membranes due to its activity increating hemifusion intermediates (44). Given this evidence,we postulate that mitofusins, through similar mechanisms thatinvolve creation of hemifusion intermediates, can induce thenucleation of holes on the OMM. In the presence of theappropriate stimulus, these holes can be employed to acceleratemitochondrial permeability transition or even participate inmitochondrial outer membrane permeabilization to allow therelease of apoptosis-inducing factors as it has been suggestedfor Drp-1 (40).

In conclusion, the present investigation shows that car-diomyocyte-specific deletion of Mfn-1 results in smaller mito-chondria that are more resistant to oxidative stress-inducedMPT and cell death. The absence of Mfn-1 did not impair leftventricular function or trigger cardiac hypertrophy. Our find-ings suggest that the endogenous levels of Mfn-1 have thecapacity to attenuate myocyte viability downstream of oxida-tive stress, an effect that could be associated with the ability ofMfn-1 to destabilize the membrane during mitochondrial fu-sion.

ACKNOWLEDGMENTS

We are grateful to Donald L. Gantz and Michael T. Kirber for assistancewith electron and confocal microscopy respectively. We are also very thankfulto Taina Rokotuiveikau and Matthew Phillippo for assistance in mousegenotyping.

GRANTS

This study was funded by National Institutes of Health Grants HL-074237(to W. C. Stanley) and HL-102874, AG-34972, AG-15052, and HL-68758 (toK. Walsh).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).

AUTHOR CONTRIBUTIONS

Author contributions: K.N.P., G.A.N., E.R.D., K.A.O., and R.F.R. per-formed experiments; K.N.P. and G.A.N. analyzed data; K.N.P. interpretedresults of experiments; K.N.P. and E.R.D. prepared figures; K.N.P. and K.W.drafted manuscript; K.N.P., E.R.D., W.C.S., and K.W. edited and revisedmanuscript; W.C.S. and K.W. conception and design of research; K.W.approved final version of manuscript.

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