Traffic 2011; 12: 972982 2011 John Wiley & Sons A/Sdoi:10.1111/j.1600-0854.2011.01215.x

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In situ Measurement of the Electrical Potential Acrossthe Lysosomal Membrane Using FRET

Mirkka Koivusalo, Benjamin E. Steinberg,David Mason and Sergio Grinstein

Program in Cell Biology, Hospital for Sick Children,Toronto, ON, Canada M5G 1X8*Corresponding author: Sergio Grinstein,sergio.grinstein@sickkids.caThese authors contributed equally to this work.

The progressive acidification of the endocytic pathwayis generated by H+ pumping of electrogenic vacuolar-type ATPases (V-ATPases) on the endosomal/lysosomalmembrane. The determinants of pH during endosomematuration are not completely understood, but the per-meability to ions that neutralize the electrogenic effectof the V-ATPase has been proposed to play a central role.If counter-ion conductance becomes limiting, the gener-ation of a large membrane potential would dominate theproton-motive force (pmf ), diminishing the pH gradientproportionally. Validation of this notion requires directmeasurement of the electrical potential that developsacross the endosomal/lysosomal membrane. To date,the measurement of lysosomal membrane potential ()in situ has been hampered by the inability to access endo-somes by electrophysiological means and the fact thatindividual organelles cannot be discerned when usingpotentiometric fluorescent dyes. Here, we describe anoninvasive procedure to estimate in intact cells,based on fluorescence resonance energy transfer (FRET).At steady state, averaged 19 mV (lumen positive)and was only partially dissipated by inhibition of theV-ATPase with concanamycin A (CcA). was consider-ably increased by alkalinization of the lysosome lumen byNH4Cl, implying that at steady state the V-ATPase oper-ates at submaximal rates and that the contribution of to pmf is relatively small. Our method should enablesystematic studies of endosomal/lysosomal potential.

Key words: electrical potential, FRET, lysosome, proton-motive force

Received 1 January 2011, revised and accepted for pub-lication 3 May 2011, uncorrected manuscript publishedonline 6 May 2011, published online 1 June 2011

During endocytosis, the internalized cargo encounters aprogressively acidic pH as it proceeds to degradativelysosomes. The proper acidification of endosomes andlysosomes plays a crucial role in the regulation of severalimportant cellular functions, including receptor-ligand dis-sociation, vesicular trafficking, hydrolysis of internalized

macromolecules and assembly of coat proteins on theorganellar surface (reviewed in 1,2). While it is clear thatthe existence of a gradual transmembrane pH gradientis central to endocytic function, the mechanism wherebythe progressive acidification is established remains largelyunknown.

The acidic pH of endosomes and lysosomes is generatedby vacuolar-type ATPases (V-ATPases), which transportprotons at the expense of ATP. Because the V-ATPase isan electrogenic pump, a transmembrane electrical poten-tial (lumen positive) tends to develop when protons aretranslocated into the compartment. The developing volt-age limits further pumping, and significant net protontranslocation requires dissipation of the electrical potentialby parallel counter-ion fluxes (24). In the presence of per-meable counter ions, the emerging pH gradient togetherwith the residual electrical potential combines to establisha passive proton-motive force (pmf ) that antagonizes theaction of the V-ATPase. A differential contribution of theelectrical and chemical components to the pmf has beensuggested as one of the mechanisms to account for theprogressive acidification of the endocytic pathway. Thus,an increasing counter-ion conductance that more effec-tively dissipates the electrical component would accountfor the greater acidification of lysosomes (3,5,6). How-ever, gradations in pH could alternatively be establishedby varying degrees of H+ leakage, which would dissipatethe forming pH gradients to different extents (3,4,6,7).

Differentiation between these models of endosome acidi-fication requires that the contribution of membrane poten-tial to the pmf can be accurately measured. There arewell-established methods for in situ pH measurement ofendocytic organelles but it has been virtually impossible tomeasure the electrical potential across membranes of theendocytic pathway. Traditionally, electrical potential acrossthe plasma membrane is measured using electrophysio-logical techniques. Measurement of membrane poten-tial across intracellular membranes by electrophysiology,however, is limited by the small size of organelles and theirinaccessibility to electrodes.Membrane-permeable poten-tiometric probes can also be used to measure potentialacross membranes. Indeed, potentiometric fluorescentdyes have been used to estimate the membrane potentialof isolated endosomes and lysosomes (3,5,6,8). However,using these dyes in intact cells is not feasible, becausethey partition across all cellular membranes, making itimpossible to resolve the fluorescence contribution of indi-vidual compartments. Here, we describe a novel methodemploying fluorescence resonance energy transfer (FRET)

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to measure in situ the steady-state membrane potentialacross the lysosomal membrane in live cells. The methodis based on the targeted delivery of a fluorescent lipidanalog to lysosomes, followed by addition of an anionic,lipid-soluble potentiometric oxonol dye.

Results

Characterization of the FRET probesWe developed two different FRET-based systems tomeasure membrane potential in lysosomes. In the firstapproach, a member of the oxonol family of potentiomet-ric dyes, bis-(1,3-dibutylbarbituric acid)trimethine oxonol(DiBAC4(3)), was selected as the FRET donor and phos-phatidylethanolamine labeled in the head group witha rhodamine derivative, L--phosphatidylethanolamine-N-(lissamine rhodamine B sulfonyl) or Rh-PE, as the FRETacceptor (Figure 1A). The uncorrected fluorescence spec-tra of the two dyes, illustrated in Figure 1B, show thatthe overlap between donor (DiBAC4(3)) emission andacceptor (Rh-PE) excitation was substantial (Figure 1B).To validate the occurrence of FRET between the dyes,

a constant amount of DiBAC4(3) was titrated with egg-phosphatidylcholine (PC) vesicles containing increasingconcentrations of Rh-PE. The emission spectrum wasrecorded using excitation of the donor at 480 nm; asillustrated in Figure 1C the peak emission of the donordecreased and acceptor emission appeared and increasedas the concentration of Rh-PE increased, consistentwith energy transfer between these two fluorophores(Figure 1C).

In the second approach, we synthesized the FRETdonor by covalently linking the coumarin derivative7-diethyaminocoumarin-3-carboxylic acid succinimidylester (DACCA-SE) to the head group of phos-phatidylethanolamine (DACCA-PE; Figure 2A), followed bypurification of the conjugate by column chromatography.The purity of the product was verified by thin-layerchromatography (TLC). Another oxonol dye, bis-(1,3-diethylthiobarbituric acid)trimethine oxonol (DiSBAC2(3)),was selected as the FRET acceptor (Figure 2A). Spec-trofluorimetric analysis of the dyes again showed a sub-stantial overlap between the donor emission and acceptorexcitation spectra (Figure 2B). To assess whether FRET

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Figure 1: Characterization of DiBAC4(3) and Rh-PE as a FRET donor and acceptor, respectively. A) The structure of DiBAC4(3) andRh-PE. B) Excitation (blue) and emission spectra (green) of DiBAC4(3) in the presence of egg-PC liposomes. Excitation (yellow) andemission spectra (red) of 0.5% Rh-PE in egg-PC liposomes. The gray bars illustrate the filter bandwidths used for visualization of FRETby microscopy. C) Emission spectra obtained in the presence of both DiBAC4(3) and Rh-PE with excitation at 480 nm. Spectra obtainedusing 300 nM DiBAC4(3) and the indicated concentrations of the acceptor (Rh-PE, 0100 nM, corresponding to 00.5% of total lipid) inegg-PC liposomes are illustrated.

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Figure 2: Characterization of DACCA-PE and DiSBAC2(3) as a FRET donor and acceptor, respectively. A) Schematic of thesynthesis of DACCA-PE; also shown is the structure of DiSBAC2(3). B) Excitation (solid purple) and emission spectra (dotted blue) ofDACCA-PE in egg-PC liposomes containing 1%DACCA-PE. Excitation (solid green) and emission spectra (dotted yellow) of DiSBAC2(3) inthe presence of egg-PC liposomes. The gray bars illustrate the filter bandwidths used for visualization of FRET by microscopy. C)Emission spectra obtained in the presence of both 200 nM (1% of total lipid) DACCA-PE in egg-PC liposomes and DiSBAC2(3) withexcitation at 400 nm; individual spectra were obtained at the acceptor (DiSBAC2(3)) concentrations indicated (0500 nM). Note that inthe absence of donor molecules there was no detectable fluorescence, even at 1 M DiSBAC2(3).

can occur between these dyes, DACCA-PE was incor-porated into egg-PC liposomes that were titrated withincreasing amounts of DiSBAC2(3), while recording theemission spectrum upon excitation of the donor at 400 nm(Figure 2C). As the amount of acceptors increased, thedonor emission progressively decreased while the accep-tor emission increased. Although not as marked as in thecase of DiBAC4(3) and Rh-PE (Figure 1C), the magnitudeof the FRET between DACCA-PE and DiSBAC2(3) wasnevertheless substantial.

Targeting FRET probes to the lysosomalcompartment in RAW264.7 macrophagesBecause the potentiometric oxonols are soluble andmem-brane permeant, they distribute throughout the cells andare unable to provide organelle-specific information. Toselectively measure lysosomal membrane potential, wetherefore used energy transfer, targeting the paired flu-orophores to the compartment of interest. In the caseof DiBAC4(3), employed as a fluorescence donor, Rh-PEwas used as an acceptor, whereas DACCA-PE was used

as a donor for DiSBAC2(3). PE derivatives with covalentlabels attached to their head-group were described ear-lier to be sorted to lysosomes after internalization (9,10).We therefore devised protocols to selectively target thefluorescent phosphatidylethanolamine derivatives to lyso-somes, using RAW264.7 cells as a model system. Thesemurine macrophages perform active constitutive endocy-tosis, expediting the labeling procedure. We used Alexa488-dextran to identify lysosomes in intact cells. A 20-hincubation with this marker ensured preferential accumu-lation in lysosomes, and residual label in endosomes wasforwarded to lysosomes during the course of the sub-sequent delivery of the labeled lipid. A fraction of thelabel may remain in late endosomes under these condi-tions, but it is likely very small and will be disregardedhereafter. After Alexa 488-dextran labeling, the cells wereincubatedwith Rh-PE complexed to BSA for 10 min on ice,to allow incorporation of the lipid into the plasma mem-brane. The label was then chased at +37C and imagedat different times. Imaging by spinning disc confocalfluorescence microscopy showed that shortly after label-ing Rh-PEwas predominantly in the plasmamembrane but

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Figure 3: Distribution of Rh-PE and DACCA-PE in live RAW264.7 macrophages. A) Cells were labeled with Rh-PE as described inMaterials and Methods and chased for the indicated times before acquiring confocal images using spinning disc microscopy. In the tworight-most panels, the lysosomes were prelabeled with Alexa 488-dextran and the localization relative to Rh-PE chased for 60 min isillustrated. B) Comparison of the localization of different organellar markers with Rh-PE chased into cells for 6090 min, as describedin Materials and Methods. The right-most panels show an enlargement of the region delineated in the adjacent overlay panels. C)Localization of DACCA-PE after a 60-min chase, compared to TMR-dextran-labeled lysosomes. Scale bars = 10 m.

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gradually moved to punctate structures which, by 60 min,colocalized with lysosomal dextran (Figure 3A). To bettercharacterize the compartment where Rh-PE resided after60 min, we compared its localization with that of variousorganellar markers (Figure 3B). At this stage, Rh-PE didnot show significant colocalization with the early endoso-mal marker Rab5-GFP or the recycling endosomal markerRab11A-GFP. Conversely, Rh-PE colocalized extensivelywith DQ-BSA, a probe that becomes fluorescent onlyin the proteolytic environment of lysosomes. Neitherthe Golgi marker galactosyltransferase-GFP (GalT-GFP),the mitochondrial marker Mitotracker, nor an endoplas-mic reticulum marker (GFP with the signal sequence ofpreprolactin and a C-terminal KDEL retention sequence)colocalized with Rh-PE, as expected (Figure 3B).

The concentration of Rh-PE attained in lysosomes usingthis procedure was estimated in parallel experimentsmea-suring the fluorescence of lysates of loaded cells preparedin PBS containing 1% Triton-X-100 and comparing themto equivalent lysates from unlabeled cells mixed withknown amounts of Rh-PE, and in parallel measuring thevolume occupied by lysosomes in the cells. The latter wasaccomplished by optical sectioning and three-dimensionalreconstruction of images of cells, where lysosomes hadbeen loaded with rhodamine-dextran (see Materials andMethods for details). Using this approach, we estimatethe concentration of Rh-PE in lysosomes to be 135 M.

To investigate DACCA-PE localization in RAW264.7 cells,this lipid analog was introduced to the plasma membraneusing methyl--cyclodextrin (mCD)-mediated transfer.After pulse labeling the cells for 5 min followed by a1-h chase, imaging by spinning disc confocal fluorescencemicroscopy showed that, as found for Rh-PE, DACCA-PEwas in punctate structures that largely colocalizedwith dextran previously chased into the lysosomalcompartment (Figure 3C). These observations indicatethat, with the labeling protocol used, both lipid analogscan be reliably and consistently targeted to lysosomes.

Imaging FRET in live RAW264.7 macrophagesTo measure FRET in macrophages, cells were first labeledwith fluorescent PE analogs followed by addition of themembrane-permeant oxonol dye to the bathing medium.Oxonols were allowed to equilibrate across all cellularmembranes and epifluorescence microscopy was used tosequentially record the emission in the donor, acceptorand FRET channels. Bleed-through of donor and acceptorfluorescences was subtracted from the FRET readingsto calculate the corrected fluorescence resonance energytransfer (cFRET; see Materials and Methods for details).Cells labeled with donor alone, whether DiBAC4(3) in thecase of the first FRET pair (Figure 4A) or DACCA-PE in thesecond case (Figure 4B), showed negligible cFRET sig-nal. Similarly, cFRET was virtually absent when only theacceptor, Rh-PE in one case (Figure 4A) or DiSBAC2(3) inthe other (Figure 4B), was present. Note also that bothpotentiometric oxonol dyes were diffusely distributed

throughout the cell, precluding visual delineation of indi-vidual organelles. Importantly, in macrophages that hadbeen labeled with both the donor and the acceptor, acFRET signal was clearly detectable and restricted to thepunctate lysosomes (Figure 4A,B).

Lysosomal membrane potential () measurementHaving confirmed the effectiveness of the FRET pairs, weproceeded to measure the lysosomal membrane potential(). The cFRET signal, a function of the oxonol concentra-tion in the immediate vicinity of the lysosomal PE probe,is an indirect measure of and must be calibrated toobtain quantitative estimates. We devised an in situ cal-ibration protocol to calculate the magnitude of fromthe cFRET values. The rationale applied to calibrationsis similar to the one used before (11) and is illustratedschematically in Figure 5A. The system consists of threecompartments: the extracellular space (compartment 1),the cytosol (compartment 2) and the intralysosomal com-partment (compartment 3). An electrical potential differ-ence PM exists across the plasma membrane, whichseparates extracellular space and cytosol, whereas a dis-tinct membrane potential is assumed to exist acrossthe lysosomal membrane. The concentration of the oxonolin each compartment (Cx) is dictated by PM and . For agiven extracellular oxonol concentration C1, the cytosolicconcentration C2 is a function of PM. The intralysosomaloxonol concentration C3, on the other hand, is dependenton both C2 and . The relationships between the fivevariables can be described by the following equations:

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We determined the value for C3 from external cali-brations performed separately for each experiment byusing DiBAC4(3) and Rh-PE as the FRET pair. To thisend, RAW264.7 cells labeled with Rh-PE were incu-bated in K+-rich buffer, which induces depolarization ofthe surface membrane. The monovalent cationophoregramicidin A was included in the medium to aid in equi-librating K+ across cell membranes and concanamycinA (CcA) to inhibit the contribution of V-ATPase to thelysosomal membrane potential. Different concentrationsof the oxonol DiBAC4(3) were added to the buffer, allowedto equilibrate across the cellular membranes and cFRETmeasured in lysosomes. Under these conditions, thepotential across both the surface and lysosomal mem-branes is assumed to be negligible and the extracellularconcentration of the oxonol should be virtually identicalto that in the cytosol and in the lysosomal lumen. Theintensity of the cFRET signal also depends on the concen-tration of Rh-PE. It is important to note that calibrations

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Figure 4: FRET imaging in liveRAW264.7 macrophages. A) Cellslabeled with only donor 300 nMDiBAC4(3) (top row), only acceptor Rh-PE (middle row), or both the donor andthe acceptor (bottom row). For eachcondition, epifluorescence microscopyimages were sequentially acquired in thedonor, acceptor and cFRET channels.The corresponding differential interfer-ence contrast (DIC) images are alsoillustrated. B) Cells labeled with onlydonor DACCA-PE (top row), only accep-tor 125 nM DiSBAC2(3) (middle row) orboth the donor and the acceptor (bot-tom row) and imaged as in (A). Scalebars = 10 m.

were always performed for each individual experimentusing cells labeled at the same time; this ensured theRh-PE (acceptor) concentration would be identical for theexperimental and calibration samples. This was validatedexperimentally by comparing the fluorescence of Rh-PE inparallel samples exposed to varying concentrations of thedonor, DiBAC4(3) (Figure S1).

The measurements obtained using increasing concen-trations of DiBAC4(3) were used to generate a calibrationcurve (Figure 5B) that enabled us to interpolateC3 from thecFRET values of experimental determinations. By usingeqn (1), we calculated C2 from the known value of C1

and using a previously determined value of 69.1 mV forPM in resting RAW264.7 cells (11). From the ratio of C3and C2 we were able to calculate the lysosomal mem-brane potential, . In otherwise untreated RAW264.7cells, the average steady-state was estimated to be18.6 1.3 mV (lumen positive; mean SE of sevenexperiments measuring at least 35 lysosomes in each;Figure 5C). Very similar results were obtained using twodifferent DiBAC4(3) concentrations, 150 or 300 nM, sup-porting the reliability of the determinations.

It is noteworthy that the calculated luminal concentrationof DiBAC4(3) (89 7.6 nM under basal conditions when

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Figure 5: Lysosomal membrane potential measurement with FRET. A) Variables for the calculation of . The system comprisesthe extracellular (compartment 1), intracellular (compartment 2) and intralysosomal (compartment 3) spaces. The concentration offree DiBAC4(3) in each compartment is indicated as Cx, where x denotes the compartment. Compartments 1 and 2 are separatedby the plasma membrane and compartments 2 and 3 by the lysosomal membrane, with transmembrane potentials PM and ,respectively. B) External calibration used to calculate C3. RAW264.7 cells were incubated with gramicidin (1 M) during the Rh-PEchase, and then transferred into K+-rich buffer containing 1 M gramicidin and 1 M CcA to collapse the membrane potentials. Differentconcentrations of DiBAC4(3) were added and the amount of cFRET was measured within the lysosomes. The amount of cFRET wasthen translated into C3 by using the calibration curve determined separately for each experiment. A representative calibration curve ofcFRET versus DiBAC4(3) concentration (C1) is shown. Data are means of 6595 lysosomes SD. C) Measurement of basal and 5 min after addition of 20 mM NH4Cl or 1 M CcA, as indicated, measured by using 300 nM or 150 nM DiBAC4(3) in the bathingmedium (data obtained at both concentrations were similar and were grouped together). Data are means of five to seven experiments SEM with at least 35 lysosomes in each. p < 0.001, p < 0.01 (ANOVA with a Dunnets test), relative to the basal condition. D)Measurement of lysosomal pH under the same conditions as in (C), obtained by ratiometric imaging of lysosomes loaded with a mixtureof FITC-dextran/OG488-dextran, as described in Materials and Methods. The inset illustrates the progressive alkalinization of lysosomesat the indicated times after addition of 1 M CcA. Data are means of three to eight experiments SEM with 10 lysosomes in each.p < 0.001 (ANOVA with a Dunnets test), relative to the basal condition. E) Representative cFRET images for each condition. Scalebar = 10 m.

the extracellular concentration was 300 nM) is compara-tively low. At face value, this amount of dye would beexpected to yield little fluorescence. However, the parti-tion coefficient of DiBAC4(3) greatly favors hydrophobicenvironments (12); therefore, the concentration of thedye within the lysosomal membrane is several orders

of magnitude higher than that in the aqueous phase ofthe lumen, clearly sufficient for detection. Indeed, thisprinciple guided our choice of Rh-PE which is located inthe bilayer as an acceptor. We tested earlier the solu-ble rhodamine-dextran as an acceptor and obtained muchweaker cFRET signals (Figure S2).

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Contribution of the V-ATPase to Next, we assessed the contribution of the electrogenicV-ATPase to , as it is the main contributor to lyso-some acidification. Proton pumping by the V-ATPasewas arrested by addition of maximally inhibitory con-centrations of the specific inhibitor CcA, and 5 min latercFRET was measured in lysosomes as above. Inhibitionof the V-ATPase decreased the cFRET values signifi-cantly (Figure 5E). Calculation of the membrane potentialrevealed a decrease of 50%, to 9.4 0.9 mV (lumenpositive; Figure 5C), indicating that the electrogenic activ-ity of the pump is an important contributor to the lysosomalmembrane potential at steady state. However, other com-ponents exist; a Donnan potential is a likely contributor tothe residual voltage. It is noteworthy that at the timewhen was measured, i.e. 5 min after addition of CcA, thechange in lysosomal pH was negligible (Figure 5D). Thelow rate of H+ (equivalent) leakage together with the largebuffering power of the lysosomes accounts for this obser-vation. Indeed, the dissipation of the transmembrane H+

gradient proceeded gradually for the following 30 min(see inset in Figure 5D). Lysosomes are rich in proteins,as evinced by their electron density, and many of theseare likely to bear a net positive charge, considering thelow prevailing pH. This would favor the generation of asizable Donnan potential.

The existence of an electrical potential together with asizable transmembrane pH gradient implies that a con-siderable pmf opposes the activity of the V-ATPase. Assuch, it is likely that the enzyme is not operating at itsfull capacity. To test this notion, we partially relieved thepmf by alkalinizing the lysosomal pH. This was accom-plished by addition of NH4+, a membrane-permeant weakbase. Independent lysosomal pH measurements verifiedthat addition of 20 mM NH4Cl to the solution bathing thecells induced a large and sustained alkalinization, frompH 4.9 to 6.6 (Figure 5D). Concomitantly, the accumula-tion of oxonol measured by cFRET increased markedly(Figure 5C,E). Calibration of this enhanced signal yieldedan average membrane potential of 28.5 1.2 mV (lumenpositive; Figure 5C). This hyperpolarization was abolishedwhen cells were (pre)treated with CcA. When both theinhibitor and NH4Cl were present simultaneously, thepotential dropped to 10 mV, indistinguishable from thatobserved in the presence of CcA alone (not shown).

Similar results were obtained when using the otherFRET pair, DACCA-PE and DiSBAC2(3): decreasedupon addition of CcA and increased with NH4Cl (notillustrated). Although producing less intense cFRET, theresults obtained with DACCA-PE and DiSBAC2(3) offerindependent confirmation of the observations made withDiBAC4(3) and Rh-PE. In addition, HeLa cells respondedsimilarly to these two treatments (not shown), indicatingthat the effects were not unique to RAW264.7 cells andthat the method is applicable to other cell types.

Discussion

To our knowledge, this is the first method described tomeasure lysosomal membrane potential in live cells. FREThad been used previously to measure the plasmalemmalmembrane potential (13,14), but intracellular organellespresent unique challenges on account of their inaccessi-bility andmuch smaller size.We estimated that lysosomesat rest have a lumen-positive of nearly 20 mV, whichis of the same order as previously suggested values forother endocytic compartments. In live cells, potentials of1020 mV were calculated from measurements of chlo-ride concentration (15), based on the premise that chloridepermeability is the dominant contributor to the endoso-mal conductance and that chloride is at electrochemicalequilibrium; these assumptions need to be validated. Themembrane potential was also measured previously in iso-lated endosomes; potentials of 2030 mV were obtainedin buffers containing Cl concentrations resembling that ofthe cytosol (6). Lastly, we recently reported that the mem-brane potential of another endocytic organelle, the phago-some, is similarly positive inside, averaging 27 mV (11). Allthese organelles share one feature in common, namelythe presence of active electrogenic H+ pumps. In the caseof lysosomes, we found that a fraction of the steady-statemembrane potential is directly attributable to the activityof the V-ATPases.

By combining themembrane potential determinations pre-sented here with independent measurements of the pHgradient (16), we can estimate directly the pmf acrossthe lysosomal membrane. In RAW264.7 cells, the lysoso-mal and cytosolic pH are 4.64 and 7.37 (16), respectively,resulting in apH of 2.73 units across the lysosomalmem-brane. From these measurements, the pmf across thelysosomal membrane was calculated to reach 18.0 kJ/molat steady state, 1.8 kJ/mol of which is contributed by theelectrical component. This antagonizing force curtails theactivity of the V-ATPase, which operates at submaximalrate in the steady state. This became apparent when thepH was reduced by addition of NH4+, which resulted ina CcA-sensitive hyperpolarization. Yet, while significant,the measured pmf is considerably lower than the the-oretical limit imposed by the energy available from ATPhydrolysis, the driving force of the V-ATPase. Assuming astoichiometry of 2 H+ per ATP hydrolysis cycle, a theoreti-cal limit of approximately 29 kJ/mol is available to generatethe lysosomal electrochemical H+ gradient (17,18). Thatthe pmf estimated empirically falls short of this valueimplies imperfect coupling. At least two mechanisms canbe readily envisaged to account for this: first, the energyconversion efficiency of the V-ATPase is

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The absolute magnitude of the membrane potential islikely to vary along the endocytic pathway. If the pmfwere to stay constant across compartments, one wouldanticipate amore positive potential at earlier stages, whichwould decline as the luminal acidification increases. Thebalance between the chemical and electrical componentsof the pmf is presumably dictated by the counter-ionpermeability. That counter ions are required for effectiveacidification has been repeatedly shown in vitro (3,5,6,8).However, such studies fail to identify the ion(s) that arephysiologically relevant. This information is best gleanedfrom measurements in situ. In live cells, a parallel Clgradient has been shown to be required for the establish-ment of a pH gradient in endosomes (15,19). In contrast,Cl is not absolutely required as a counter ion to acidifylysosomes in situ; efflux of luminal cations can equally sus-tain lysosome acidification (16). The additional counter-ionpathway(s) possibly enable the lysosome to more readilycompensate the electrogenicity of the V-ATPase, enablingit to acidify the lysosomal lumen more profoundly.

While having provided the first direct measurements oflysosomal membrane potential, we regard the main valueof this report as being technical in nature. We believe thatthe FRET-based assay introduced here to determine lyso-somal membrane potential can be modified and extendedto measure the membrane potential of other endocyticand secretory compartments that to date have remainedinaccessible by other approaches. A variant of this proce-dure could be used to test the role of the electrogenicNa,K-ATPase in early endosomes, where it purportedlylimits acidification by increasing membrane potential (20).Similarly, the operation of other ion channels, transportersand pumps residing within the endocytic pathway rep-resented could be similarly probed. Such measurementswould improve our understanding of ion transport and pHhomeostasis in endomembranes.

Materials and Methods

ReagentsDACCA-SE, DiBAC4(3), DiSBAC2(3), DQ-BSA green, Mitotracker green,nigericin and tetramethylrhodamine (TMR)-, rhodamine B-, Alexa 488-, flu-orescein isothiocyanate (FITC)- and Oregon Green 488 (OG488)-conjugateddextrans [all of molecular weight (MW) 10 000] were from Invitro-gen. Egg-Rh-PE, egg-PC and 16:0/18:1-PE [palmitoyloleoylphosphatidylethanolamine (POPE)] were from Avanti Polar Lipids. FugeneHD was fromRoche. All other reagents were from Sigma-Aldrich.

Isotonic Na+-rich buffer contained 140 mM NaCl, 3 mM KCl, 1 mM MgCl2,1 mM CaCl2, 5 mM glucose and 20 mM HEPES (pH 7.4). In K+-rich buffer,NaCl was replaced by 100 mM K-glutamate and 43 mM KCl.

DACCA-PE synthesisPOPE in CHCl3 was combined with 5.5 molar excess of DACCA-SEin dimethyl sulfoxide, and the mixture incubated overnight with vigorousshaking. The reaction mixture was dried under N2, resuspended in 98:2CHCl3:MeOH and layered onto a 2-cm silica column in the same solvent.The column was washed with 98:2 CHCl3:MeOH until all the unreactedDACCA-SE had been washed out (5 2 mL). The DACCA-PE was eluted

out in consecutive 90:10:0.5 CHCl3:MeOH:H2O fractions monitored byTLC. Fractions containing the end product were used for cell labeling.

FRET pair characterizationTo measure the excitation and emission spectra of Rh-PE and DACCA-PEin the absence and presence of the oxonols, these lipids were incorpo-rated into egg-PC vesicles at different concentrations. Lipids were driedunder N2 and reconstituted into multilamellar vesicles by vortexing to20 M final lipid concentration in PBS. Excitation and emission spectraof DiBAC4(3) and DiSBAC2(3) were measured in the presence of egg-PCliposomes without the fluorescent lipids. Spectra were acquired by usingan F-2500 Fluorescence Spectrophotometer (Hitachi).

Cell culture, plasmids and transfectionsRAW264.7 macrophages (ATCC number TIB-71) were obtained from theAmerican Type Culture Collection and grown at 37C under 5% CO2 inRPMI-1640 supplemented with 5% FBS (Wisent). The plasmids encod-ing Rab5A-GFP (21) and Rab11A-GFP (22) have been described elsewhereand were a kind gift from Dr John Brumell (Hospital for Sick Children,Toronto). Preprolactin signal sequence-GFP-KDEL has also been previouslydescribed (23). GalT-GFP was from Addgene (plasmid #11929) (24). Trans-fections were performed with FugeneHD following the manufacturersinstructions.

Labeling of cells with DACCA-PEDACCA-PE was introduced into RAW264.7 cells using mCD-mediatedtransfer (25,26). For labeling one 18-mm coverslip, 36 g DACCA-PEand 5 g cholesterol were dried under N2; 100 mM mCD was addedand the mixture probe sonicated for 3 2 min (0.9 g/L DACCA-PE and0.125 g/L cholesterol in the mixture). Forty microliters of the mixturewas added to cells in 500-L serum-free RPMI for a final concentrationof 67 and 9.3 g/mL for DACCA-PE and cholesterol, respectively, and7.4 mM for mCD. The cells were pulsed with the lipid mixture for 5 minat 37C, washed and chased for 15 min in serum-free RPMI followed bya 1560-min chase in RPMI + 5% FBS to remove any residual DACCA-PEleft on the plasma membrane. Cells were then transferred to serum-freeHEPES-buffered RPMI (HPMI) or Na+-rich buffer for imaging.

Labeling of cells with Rh-PERh-PE was freshly complexed with fatty acid-free BSA by adding 25 Lof 1 mg/mL Rh-PE solution in 1:1 CHCl3/MeOH dropwise into 4 mL of0.5 mg/mL BSA in PBS while vigorously vortexing for 12 min. A gentlestream of N2 was blown on the complex before placing it on ice. RAW264.7cells grown on 18- or 25-mm coverslips were washed with ice-cold PBS,and incubated with the Rh-PEBSA complex for 10 min on ice to firstintroduce the lipid to the plasma membrane. Cells were washed withwarm HPMI or Na+-rich buffer, and chased in this buffer at 37C for6090 min to label the lysosomes.

Lysosomal membrane potential measurementsA 150 or 300 nM DiBAC4(3) was added into the bathing medium of Rh-PE-labeled cells and cells allowed to equilibrate for 5 min at 37C. ForDACCA-PE-labeled cells, 125 nM DiSBAC2(3) was added to the bathingmedium and cells equilibrated as above. Fifteen to 30 baseline images ofdifferent fields of the donor, acceptor and FRET channels were acquired.The cells were then treated with 1 M CcA or 20 mM NH4Cl, or both. After5 min of incubation, 1530 images in the three channels were acquired.For the determination of the donor and acceptor bleed-through coefficients,samples with donor and acceptor only were imaged in each experiment.

To determine the amount of DiBAC4(3) within the lysosomes, separatecalibration samples were treated as follows. During chase with Rh-PE,1 M gramicidin was included in HPMI. Cells were then washed withK+-rich buffer and incubated with 25300 nM DiBAC4(3), 1 M CcA and

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1 M gramicidin A in K+-rich buffer to collapse the membrane potential.After 5 min of incubation, 1530 images of different fields in all thethree channels were acquired. In this way, a calibration curve relatingcFRET to DiBAC4(3) concentration was constructed, and the intralysoso-mal DiBAC4(3) concentration determined from the linear fit of the data.To adjust the microscope settings for each experiment, measurementswere started with the highest DiBAC4(3) concentration in K+-rich buffer,which gave the highest DiBAC4(3) fluorescence inside the cells. wascalculated as described in Results.

Live-cell imaging and FRETCoverslips were mounted in Chamlid imaging chambers (Live Cell Instru-ment Inc.) and maintained at 37C on the stage of a Leica DM IRB micro-scope equipped with filter wheels (Sutter Instruments) to independentlyalternate between different excitation and emission filters. Light froman EXFO X-Cite 120 lamp (Exfo Life Sciences Group) was directed tothe sample by using a dichroic mirror. Emitted light was captured bya Cascade II CCD camera (Photometrics). The filter wheel and cam-era were under the control of METAMORPH/METAFLUOR software (MolecularDevices). In FRET experiments, three fluorescent channels were seriallyacquired. The filter set configurations (listed as excitation/dichroic/emissionfilters, all in nanometers bandwidth) used in the FRET experimentswere as follows: donor DiBAC4(3) 475 60/495/510 40, acceptor Rh-PE543 22/565/580 25 and FRET 475 60/565/580 25; donor DACCA-PE417 60/425/470 40, acceptor DiSBAC2(3) 543 22/565/580 25 andFRET 417 60/565/580 25. A PL Fluotar 100/numerical aperture (NA)1.3 oil immersion objective was used. cFRET was calculated using themethod of Youvan et al. (27). The donor and acceptor bleed-through coeffi-cients were determined by acquiring images with donor or acceptor alone,respectively, in independent samples. All microscope parameters, includ-ing exposure times and camera gains, were kept constant across eachexperiment. Fluorescence intensity measurements were performed usingIMAGEJ software. Images were background-subtracted before analysis.

Lysosomal pH measurementspH measurements were performed by ratiometric imaging of pH-sensitivedextrans targeted to lysosomes using a pulse-chase protocol. RAW264.7cells were incubated in serum-free growth medium for 30 min and thenlabeled with a mixture of 0.05 mg/mL OG488-dextran and 0.25 mg/mLFITC-dextran for 620 h at 37C. This combination of dyes with disparatepKa values was selected to cover a wide range of pH. Cells were subse-quently washed, chased under serum-free conditions for 30 min to 1 h andimaged with the microscopy system described for FRET, using alternateexcitation through 485 10-nm and 438 12-nm filters, a 505-nm dichroicmirror and a 535 20-nm emission filter. At the end of each experiment,an in situ calibration was performed by bathing the cells in K+-rich solutioncontaining 20 mM HEPES, 2-(N-morpholino)ethanesulfonic acid (MES) oracetate, as appropriate, plus 10 g/mL nigericin and buffered to a pH rangeof 47, as described (16).

Labeling of cells with other fluorescent markersTo label lysosomes with Alexa 488-dextran or TMR-dextran for localization,RAW264.7 cells were incubated in serum-free growth medium for 30 minand then labeled with 0.10.25 mg/mL dextran for 20 h at 37C. Afterextensive washing, cells were labeled with Rh-PE or DACCA-PE andchased as described above. To label lysosomes with green DQ-BSA, cellswere incubated with 50 g/mL DQ-BSA for 1 h at 37C before labelingwith the fluorescent lipids. To label mitochondria, RAW264.7 cells weretreated with 500 nM Mitotracker green.

Spinning disc confocal fluorescence microscopyTo assess the detailed localization of Rh-PE and DACCA-PE in the cellsrelative to organellar markers, images were acquired using a QuorumWaveX (Guelph) spinning disc confocal microscopy system based on aZeiss Axiovert 200M microscope. Cells were maintained at 37C using

a Live Cell Instrument environmental control system. The samples wereexcited with diode-pumped solid-state laser lines (Applied Research) usinga 491-nm laser line and 520-nm emission filter for green fluorophores,a 561-nm laser line and 590-nm emission filter for Rh-PE and a 405-nmlaser line and 457-nm emission filter for DACCA-PE. A 63/NA 1.4 oilimmersion objective was used. Images were acquired using a HamamatsuC9100-13 ImagEM back-thinned electron multiplier CCD camera driven bythe VOLOCITY 4.1.1 software (Perkin Elmer).

Estimation of intralysosomal Rh-PE concentrationCells were loaded with Rh-PE as described above and after a 1-h chasethey were scraped and suspended into PBS containing 1% Triton-X-100. The fluorescence emitted by Rh-PE was measured using a Hitachispectrofluorimeter. The amount of Rh-PE per cell was estimated byinterpolation in a standard calibration curve obtained adding knownamounts of Rh-PE into a cell lysate prepared from unlabeled cells; theamount of unlabeled lysate used in this instance was identical to thatused for the loaded cells above. To estimate the volume of lysosomes,RAW264.7 cells were labeled with 0.2 mg/mL rhodamine B-dextran for16 h, chased for 1 h and imaged by spinning disc confocal microscopy.Serial optical slices were collected and used to construct a three-dimensional image. The average volume occupied by the lysosomes ineach cell was estimated by using the IMAGEJ (NIH) 3Dobject counter plug-in.

Acknowledgments

This work was supported by grants from the Canadian Institutes ofHealth Research (CIHR grant MOP4665) and the Canadian Cystic FibrosisFoundation. M. K. was supported by the Ella and Georg EhrnroothFoundation. B. E. S. was the recipient of studentships from theMacLaughlin Centre for Molecular Medicine and the CIHR. S. G. holdsthe Pitblado Chair in Cell Biology and is cross-appointed to the Departmentof Biochemistry of the University of Toronto.

Supporting Information

Additional Supporting Information may be found in the online version ofthis article:

Figure S1: Fluorescence intensities of Rh-PE and DiBAC4(3) inexternal calibration samples. Rh-PE-labeled RAW264.7 cells wereincubated in K+-rich buffer containing 1 M gramicidin, 1 M CcAand the indicated concentrations of DiBAC4(3). Background-subtractedfluorescence intensities of Rh-PE and DIBAC4(3) in cells from 13 to 19imaged fields from a representative experiment are shown. Data aremeans SD.

Figure S2: Comparison of FRET imaging using DiBAC4(3) as the donorand either fluid-phase rhodamine B-dextran or membrane-boundRh-PE as the acceptor. The images were acquired the same day inparallel experiments using the same cell culture and the same acquisitionsettings for each individual channel to make the intensity measurementscomparable. Scale bar = 10 m.

Please note: Wiley-Blackwell are not responsible for the content orfunctionality of any supporting materials supplied by the authors.Any queries (other than missing material) should be directed to thecorresponding author for the article.

References

1. Marshansky V, Futai M. The V-type H+-ATPase in vesicular traf-ficking: targeting, regulation and function. Curr Opin Cell Biol2008;20:415426.

2. Weisz OA. Acidification and protein traffic. Int Rev Cytol 2003;226:259319.

Traffic 2011; 12: 972982 981

Koivusalo et al.

3. Fuchs R, Male P, Mellman I. Acidification and ion permeabilities ofhighly purified rat liver endosomes. J Biol Chem1989;264:22122220.

4. Grabe M, Oster G. Regulation of organelle acidity. J Gen Physiol2001;117:329344.

5. Van Dyke RW. Acidification of rat liver lysosomes: quantitation andcomparison with endosomes. Am J Physiol 1993;265:C901C917.

6. Van Dyke RW, Belcher JD. Acidification of three types of liverendocytic vesicles: similarities and differences. Am J Physiol1994;266:C81C94.

7. Lukacs GL, Rotstein OD, Grinstein S. Determinants of the phagoso-mal pH in macrophages. In situ assessment of vacuolar H(+)-ATPaseactivity, counterion conductance, and H+ leak. J Biol Chem1991;266:2454024548.

8. Van Dyke RW, Scharschmidt BF, Steer CJ. ATP-dependent protontransport by isolated brain clathrin-coated vesicles. Role of clathrinand other determinants of acidification. Biochim Biophys Acta1985;812:423436.

9. Verkade HJ, Zaal KJ, Derksen JT, Vonk RJ, Hoekstra D, Kuipers F,Scherphof GL. Processing of the phospholipid analogue phosphatidyl(N-sulphorhodamine B sulphonyl)ethanolamine by rat hepatocytes invitro and in vivo. Biochem J 1992;284:259265.

10. Willem J, ter Beest M, Scherphof G, Hoekstra D. A non-exchangeablefluorescent phospholipid analog as a membrane traffic marker of theendocytic pathway. Eur J Cell Biol 1990;53:173184.

11. Steinberg BE, Touret N, Vargas-Caballero M, Grinstein S. In situmeasurement of the electrical potential across the phagosomalmembrane using FRET and its contribution to the proton-motiveforce. Proc Natl Acad Sci U S A 2007;104:95239528.

12. Zeng Y, Han X, Gross RW. Phospholipid-subclass-specific partition-ing of lipophilic ions in membrane-water systems. Biochem J1999;338:651658.

13. Chanda B, Blunck R, Faria LC, Schweizer FE, Mody I, Bezanilla F. Ahybrid approach to measuring electrical activity in genetically specifiedneurons. Nat Neurosci 2005;8:16191626.

14. Gonzalez JE, Tsien RY. Improved indicators of cell membranepotential that use fluorescence resonance energy transfer. ChemBiol 1997;4:269277.

15. Sonawane ND, Thiagarajah JR, Verkman AS. Chloride concentrationin endosomes measured using a ratioable fluorescent Cl indicator:

evidence for chloride accumulation during acidification. J Biol Chem2002;277:55065513.

16. Steinberg BE,Huynh KK, Brodovitch A, Jabs S, Stauber T, Jentsch TJ,Grinstein S. A cation counterflux supports lysosomal acidification.J Cell Biol 2010;189:11711186.

17. Kettner C, Bertl A, Obermeyer G, Slayman C, Bihler H. Electrophysi-ological analysis of the yeast V-type proton pump: variable couplingratio and proton shunt. Biophys J 2003;85:37303738.

18. Tomashek JJ, Brusilow WS. Stoichiometry of energy coupling byproton-translocating ATPases: a history of variability. J BioenergBiomembr 2000;32:493500.

19. Sonawane ND, Verkman AS. Determinants of [Cl] in recycling andlate endosomes and Golgi complex measured using fluorescentligands. J Cell Biol 2003;160:11291138.

20. Fuchs R, Schmid S, Mellman I. A possible role for Na+,K+-ATPase inregulating ATP-dependent endosome acidification. Proc Natl Acad SciU S A 1989;86:539543.

21. Scott CC, Cuellar-Mata P, Matsuo T, Davidson HW, Grinstein S. Roleof 3-phosphoinositides in the maturation of Salmonella-containingvacuoles within host cells. J Biol Chem 2002;277:1277012776.

22. Rzomp KA, Scholtes LD, Briggs BJ, Whittaker GR, Scidmore MA.Rab GTPases are recruited to chlamydial inclusions in both aspecies-dependent and species-independent manner. Infect Immun2003;71:58555870.

23. Dayel MJ, Hom EF, Verkman AS. Diffusion of green fluorescentprotein in the aqueous-phase lumen of endoplasmic reticulum.Biophys J 1999;76:28432851.

24. Cole NB, Smith CL, Sciaky N, Terasaki M, Edidin M, Lippincott-Schwartz J. Diffusional mobility of Golgi proteins in membranes ofliving cells. Science 1996;273:797801.

25. Kainu V, Hermansson M, Somerharju P. Introduction of phospholipidsto cultured cells with cyclodextrin. J Lipid Res 2010;51:35333541.

26. Koivusalo M, Jansen M, Somerharju P, Ikonen E. Endocytic traffickingof sphingomyelin depends on its acyl chain length. Mol Biol Cell2007;18:51135123.

27. Youvan DC, Silva CM, Bylina EJ, Coleman WJ, Dilworth MR, YangMM. Calibration of fluorescence resonance energy transfer inmicroscopy using genetically engineered GFP derivatives on nickelchelating beads. Biotechnology et alia 1997;3:118.

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