Temporal Analysis of the Nuclear-to-cytoplasmic Translocation of a Herpes Simplex Virus 1 Protein by Immunofluorescent Confocal Microscopy

This protocol has been used to study the nuclear-to-cytoplasmic translocation of HSV-1 ICP0. ICP0 undergoes subcellular trafficking during HSV-1 infection (Figure 1). Likely, ICP0 interacts with various cell pathways to carry out different functions at different locations. This enables ICP0 to fine tune its multiple functions in the tug-of-war with human host¹³. However, how ICP0 coordinates the multiple functions in a spatial-temporal manner has not been well studied. With the fluorescent microscopy protocol described above, we started to analyze the molecular basis of ICP0 nuclear-to-cytoplasmic translocation. As of now, we have identified the ICP0 C-terminal 35 amino acids as a required element important for this translocation. In the absence of C-terminus, ICP0 is restrained within the nucleus throughout infection (Figure 3). We have also found that an ICP0 E3 ligase-dependent nuclear retention force delays the nuclear-to-cytoplasmic translocation in U2OS cells¹². Furthermore, we have discovered that ICP0 C-terminus and the expression of late viral proteins cooperate to overcome the nuclear retention and facilitate cytoplasmic translocation¹². Currently, we are using this protocol to screen for the late viral proteins involved in the ICP0 nuclear-to-cytoplasmic translocation.

The protocol was initially developed to study the dynamic trafficking of ICP0 in HSV-1 infection. As shown in Figure 1, early in HSV-1 infection, ICP0 is colocalized with ND10, where several key components of cellular restrictive factors and ND10 components such as PML and Sp100⁸, are degraded. After degrading ND10 key constituents, ICP0 diffuses throughout the nucleus and late in infection, ICP0 is translocated to the cytoplasm. Because ICP0 undergoes de novo synthesis upon infection, the initial protein abundancy is very low and then a robust viral synthesis will quickly obscure the movement of any individual molecules, which makes it difficult to track a single molecule using live imaging technology. Therefore, we deliberately chose not to use live imaging. Instead, we adopted the above protocol to study the steady state ICP0 localization at different infection points, which served us well in tracking ICP0 temporal movement in a population of HSV-1 infected cells.

For a high signal-to-background ratio in confocal analysis, two critical steps are noteworthy in the wet-bench part of this protocol. First, the 4-well staggered slides allow multiple samples to be handled on one single slide. It greatly saves the usage of precious reagents like viruses and antibodies. However, because the volume held in each well is so small, residual buffer not completely cleared during buffer changes can interfere with the subsequent reagent. Therefore, in each buffer switch, a thorough aspiration is needed before adding the new buffer. Second, based on our experiences, the extent of cell crosslinking and membrane permeability is important for the clarity of fluorescence signals. We have set an empirical number of 10 min for both paraformaldehyde and nonionic surfactant treatments. We found that time much longer or shorter than 10 min can decrease the signal-to-background ratio. As shown in Figure 1 and Figure 2, as well as in a previous study¹², images obtained in our experiments are crystal clear. The prominent blue signal that clearly outlines the nuclear boundary is key to determining the subcellular distribution of ICP0. In the computational part of this protocol, one crucial step is to set a constant threshold to eliminate the background. A successful staining with high signal-to-background ratio is the key to a lower threshold line and better signal contrast. Keeping a constant threshold for all samples in the same experiment, however, is the foundation for the quantitative documentation of ICP0 (Figure 2 and Figure 3).

The protocol can also serve as a general tool to study subcellular trafficking for other viral or cellular proteins when a suitable live imaging method is lacking. In live imaging technique, cells are kept at optimal physiological environment to maintain cell metabolic status^14,15. A basic requirement for live cell imaging is to fluorescently label the target protein, which can be achieved by fusing the target protein with a fluorescent tag¹⁶, or to deliver a fluorophore conjugated molecule specific for the target protein¹⁷. In either case, problems may rise if the fusion of fluorescent tag changes target protein property or fluorophore conjugated molecule has difficulty to cross cell membrane. Photobleaching that causes cell damage in the process is an additional concern in live imaging¹⁸. Therefore, new strategies to overcome the limitation of live imaging continue to be the frontier of technology development. The protocol we described here provides temporal analysis of the steady state confocal images, which can serve as an alternative tool when a proper live imaging method is unavailable. The method is easy and reliable. It provides clear detection of protein subcellular localization with minimum background. Using confocal software, we are able to quantitatively analyze the percentage of cells with different distributions of the target protein in a cell population and document the movement of target protein at different cell phases.