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1Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, 2Howard Hughes Medical Institute, Harvard Medical School
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We present an in vitro, two-color fluorescence assay to visualize the fusion of single virus particles with a fluid target bilayer. By labeling viral particles with fluorophores that differentially stain the viral membrane and its interior, we are able to monitor the kinetics of hemifusion and pore formation.
Floyd, D. L., Harrison, S. C., van Oijen, A. M. Method for Measurement of Viral Fusion Kinetics at the Single Particle Level. J. Vis. Exp. (31), e1484, doi:10.3791/1484 (2009).
Membrane fusion is an essential step during entry of enveloped viruses into cells. Conventional fusion assays typically report on a large number of fusion events, making it difficult to quantitatively analyze the sequence of the molecular steps involved. We have developed an in vitro, two-color fluorescence assay to monitor kinetics of single virus particles fusing with a target bilayer on an essentially fluid support.
Influenza viral particles are incubated with a green lipophilic fluorophore to stain the membrane and a red hydrophilic fluorophore to stain the viral interior. We deposit a ganglioside-containing lipid bilayer on the dextran-functionilized glass surface of a flow cell, incubate the viral particles on the planar bilayer and image the fluorescence of a 100 x 100 μm2 area, containing several hundreds of particles, on a CCD camera. By imaging both the red and green fluorescence, we can simultaneously monitor the behavior of the membrane dye (green) and the aqueous content (red) of the particles.
Upon lowering the pH to a value below the fusion pH, the particles will fuse with the membrane. Hemifusion, the merging of the outer leaflet of the viral membrane with the outer leaflet of the target membrane, will be visible as a sudden change in the green fluorescence of a particle. Upon the subsequent fusion of the two remaining distal leaflets a pore will be formed and the red-emitting fluorophore in the viral particle will be released under the target membrane. This event will give rise to a decrease of the red fluorescence of individual particles. Finally, the integrated fluorescence from a pH-sensitive fluorophore that is embedded in the target membrane reports on the exact time of the pH drop.
From the three fluorescence-time traces, all the important events (pH drop, lipid mixing upon hemifusion, content mixing upon pore formation) can now be extracted in a straightforward manner and for every particle individually. By collecting the elapsed times for the various transitions for many individual particles in histograms, we can determine the lifetimes of the corresponding intermediates. Even hidden intermediates that do not have a direct fluorescent observable can be visualized directly from these histograms.
Glass cover slip functionalization
The planar bilayer used in the fusion assay is supported on a hydrated film of dextran. Dextran acts as a spacer between the planar bilayer and glass surface. This prevents membrane components from becoming stuck on the glass surface and also provides space in to which the contents of a virus particle can escape upon fusion. Glass coverslips are functionalized through treatment with an epoxy silane, which allows us to chemically bond dextran to the glass (Elender, et al. 1996).
Supported lipid bilayers are formed by adsorbing liposomes to a dextran functionalized coverslip. The adsorbed liposomes fuse with eachother until membrane ruptures and spreads flat over the surface.
Microfluidic flow cell preparation
A simple microfluidic flow cell is made by sandwiching double stick tape between a quartz slide and a functionalized coverslip.
Virus particle labeling
The experiment is performed on a fluorescence microscope equipped with a high NA oil immersion objective suitable for total internal reflection fluorescence microscopy. The 488 and 568 nm lines from an argon/krypton gas laser are used to excite the Rh110C18 and SRB labeled virus. Simultaneous imaging of each label is accomplished by splitting the green and red fluorescence emission with a dichroic mirror and independently focusing the images onto separate halves of a electron multiplying CCD camera.
Performing the Assay
Execution of the fusion assay involves a stepwise assembly of the biochemical components within the flow cell: assembly of the planar lipid bilayer, docking of labeled virus particles, coating the surface with fluorescein, and initiation of the reaction with low pH buffer (Figure 1A).
Figure 1B shows snapshots of bilayer docked virions before and during fusion. Each diffraction limited spot represents an individual virus particle. Red and green fluorescence have been separately imaged onto the top and bottom halves of a CCD camera, allowing simultaneous observation of the viral envelope and content labels. There are typically twice as many spots in the green channel compared to the red, and this reflects the lower efficiency of labeling by the content dye compared to the envelope label. The green background fluorescence emitted from surface bound fluorescein disappears upon arrival of the citrate buffer and allows determination of the start time of the fusion reaction. Hemifusion events are detected as bursts of green fluorescence as the quenched Rh110C18 in the viral envelope diffuses across the hemifusion stalk and is diluted in the planar membrane. An outward expanding fluorescent cloud can be seen around hemifused particles, demonstrating free diffusion of the fatty-acyl linked dye in the planar membrane. Pore formation is observed as the decay of red fluorescence from SRB trapped within the interior of the viral particles. The opening of a fusion pore allows the dye to escape into the aqueous space below the planar bilayer and out of the observation region.
Fluorescence intensity trajectories for each particle, plotted from the integrated intensities of individual particles, facilitate determination of the yield and times of hemifusion and pore formation (Fig. 1C). Hemifusion and pore formation event times are determined as the point at which the maximum slope of a Rh110C18 fluorescence burst or SRB signal decay occurs. In a successful experiment, about 50 percent of the particles labeled with Rh110C18 yield dequenching signals, and 30 percent of the SRB labeled particles give useable signal. Of the particles labeled with both dyes, approximately 10 percent show both lipid mixing and pore formation signals.
Fig. 2A-C shows the distribution of hemifusion and pore formation lag times from a typical experiment. Fig.2D-F shows event distributions of events combined from five experiments performed under the same conditions. Even with a modest number of observations, the shape of the histograms is apparent. Because each experiment is synchronized by detection of the pH drop time, multiple experiments can be combined and detailed information about rate limiting intermediate steps can be obtained.
Figure 1. Experimental design. A) Virus particles are labeled with two fluorescent dyes to monitor the kinetics of hemifusion and fusion pore formation. Fluorescence is collected by a high-NA microscope objective and imaged onto a CCD. B) Fluorescence images before (left) and during (right) the fusion of individual viral particles. Top and bottom half of each image correspond to the red and green fluorescence, respectively, of the same ~ 50 x 100 mm2 area of the supported bilayer. C) The fluorescence intensity of the red SRB viral content tracer (upper trace), the green Rh110C18 membrane dye (middle trace), and the fluorescein pH sensor (lower trace) provide exact times elapsed between pH drop, hemifusion, and fusion. Please click here to see a larger version of figure 1.
Figure 2. Fusion kinetics of fluorescently labeled virus. Distributions of the time elapsed between the pH drop and hemifusion (A, D) and pore formation (B,E). The rise and decay of the distributions indicate the presence of multiple intermediate steps. The transition between hemifusion and opening of a fusion pore for individual particles is exponentially distributed, suggesting a single rate-limiting step. Panels A-C show the results of a single experiment. Panels D-F are compiled from five separate experiments performed under the same conditions. Please click here to see a larger version of figure 2.
Preparation of fluid and continuous supported lipid bilayers can be challenging. Trace amounts of contaminating material or surface defects will prevent spreading of bilayers. Careful cleaning and deposition of a uniform layer of dextran are essential.
Prolonged imaging of virus particles can bleach the fluorescent labels or cause them to become inactivated. Photo-damage of this kind is well known in the bio-imaging and single molecule fields and is generally believed to stem from the generation of reactive oxygen species by the excited fluorophores To minimize photo-damage, we generally minimize the excitation laser power and avoid prolonged exposure prior to starting the experiment. Depletion of oxygen from the buffers with one of the several oxygen scavenging systems reported may prove useful.
The ability to get information about transient intermediate states requires that the fusion reaction be precisely synchronized. The flow rate used and the dead volume of the inlet tubing and flow cell channel are factors that can limit synchronization. Fusion is initiated by exchanging neutral for acidic pH buffer. However, as the acidic buffer travels through the inlet tubing into the flow cell, the interface between the two buffers mixes through diffusion and becomes progressively less well defined. The resulting pH gradient at the bilayer surface tends to blur determination of the start time. Furthermore, since fusion kinetics is dependent on proton concentration, a pH gradient at the start of the experiment could complicate interpretation of the results. In the experiment demonstrated here, the kinetics of fusion is much slower than the pH transition time, and the gradient effect does not significantly affect fusion kinetics. We are currently working on modifications to our flow cells that would allow us to prime the inlet tubing with the desired buffer before diverting the flow into the flow cell.
The work was supported by NIH grants AI57159 (to S.C.H.) and AI72346 (to A.M.v.O.). S.C.H. is a Howard Hughes Medical Institute Investigator.
|Corning cover glass squares, 25 mm||Sigma-Aldrich||CLS286525|
|Fused silica slides||Technical Glass|
|Staining rack||Thomas Scientific||8542E40|
|(3- glycidoxypropyl)trimethoxysilane||Gelest Inc.||SIG5840.1|
|Dextran T-500||Pharmacosmos||5510 0500 4006|
|1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC)||Avanti Polar Lipid, Inc||850375|
|1-palmitoyl-2oleoyl-sn-glycero-3-phosphocholine (POPC)||Avanti Polar Lipid, Inc||850457|
|Cholesterol||Avanti Polar Lipid, Inc||700000|
|N-((6-(biotinoyl)amino)hexanoyl)-1,2-dihexadecanoyl-sn-glycero-3-phosph–thanolamine, triethylammonium salt (biotin-X DHPE)||Invitrogen||B1616|
|Streptavidin, fluorescein conjugate||Invitrogen||S-869|
|Disialoganglioside GD1a from bovine brain||Sigma-Aldrich||G2392|
|Mini Extruder||Avanti Polar Lipid, Inc||610000|
|0.1 micron polycarbonate membrane filters||Whatman, GE Healthcare||800309|
|10 mm drain discs (membrane supports)||Whatman, GE Healthcare||230300|
|Sulforhodamine b sodium salt||S1402|
|Octadecyl rhodamine 110 (Rh110C18)||See Floyd et al. 2008 for synthesis protocol|
|PD-10 desalting columns||GE Healthcare||17-0851-01|
|Nikon fluorescence microscope||Nikon Instruments||TE2000-U|
|Plan Apo TIRF 60x 1.45 NA objective||Nikon Instruments|
|Innova 70C Spectrum Ar/Kr laser||Coherent Inc.|
|Syringe Pump||Harvard Apparatus||702213|
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