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Epifluorescent microscopy to evaluate bacteriophage capsid integrity
 
Kerusha Naicker and Steven I. Durbach
University of the Witwatersrand, Johannesburg, Gauteng, South Africa
BioTechniques, Vol. 43, No. 4, October 2007, pp. 473–476
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Abstract

The standard method to evaluate capsid integrity of DNA-based viruses, which depends on access to DNase I, relies on the removal of the capsid by solvent extraction and then the evaluation of the nucleic acid products by electrophoresis. Our method, which is based on the direct detection of capsid-borne DNA directly through the use of epifluorescent microscopy, negates the requirement of DNA extraction.

Virus capsid integrity can be measured indirectly by infectivity studies (1). A more informative method assesses capsid integrity by evaluating inaccessibility of DNase I to virus DNA. The efficacy of the DNase I treatment is evaluated by gel electrophoretic analysis of the extracted nucleic acid (2,3).

In recent years, staining environmental samples with asymmetrical cyanine dyes (4), such as SYBR® Green I, and subsequent visualization with epifluorescent microscopy has become the standard approach for the detection of dsDNA bacteriophages and viruses (5,6). The principle of this method is that the SYBR Green I dye is able to penetrate bacteriophage capsids and intercalate in the nucleic acid, displaying a significantly higher affinity for dsDNA than other nucleic acid types (7,8). The dsDNA is concentrated within the capsid and accordingly is detectable as dots under 1000× magnification using epifluorescent microscopy (5).

Given the robustness of this method, we reasoned that it could be adapted to a simple assay for capsid integrity with the advantage of circumventing solvent-mediated disruption of the capsids following destabilizing treatments, which does not depend on solvent sensitivity of capsid proteins. This will allow an in situ confirmation of DNase I sensitivity. We used epifluorescent microscopy to analyze destabilized T4 virions for DNase I accessibility by their ability to be stained by SYBR I Green dye.

Capsids were destabilized by heat treatment of CsCl-purified T4 phage (9). This was done by dispensing 1 mL T4 [109 plaque forming units (pfu)] contained in phage buffer (10 mM Tris, pH 8.0, 10 mM MgSO4, 68 mM NaCl, 10 mM CaCl2) into glass McCartney bottles and boiling (98°C) for 20 min. Plaque overlays showed undetectable viability following this treatment (data not shown). Transmission electron microscopy (TEM) was done to assess the impact of the heat treatment on capsid structure. Approximately 5 µL sample (titer = 1 × 106 pfu/mL) were placed on carbon-coated Formvar grids for 20 min. The excess fluid was gently drawn off with filter paper. Five microliters of stain (1% uranyl acetate in 45% ethanol) were then placed on the grid for 3 min, and excess fluid was drawn off with filter paper. The grids were washed with double deionized water, and the excess fluid drawn off. The grids were then dried for 48 h at room temperature before being viewed under the transmission electron microscope (JEOL-JEM-100S, Japan). TEM confirmed capsid destabilization (Figure 1B) compared with the untreated T4 phage control (Figure 1A) in that the heat-treated capsids appeared deformed yet intact. We then investigated whether the heat-treated capsids were detectable by epifluorescent microscopy essentially as described by Noble and Fuhrman (5). although rather than environmental material, 1–5 mL CsCl-purified T4 (titer: 1 × 106 pfu/mL) were filtered through a 0.02-µM Anodisc® 25 filter (Whatman, Kent, UK). The staining method followed was also as described by Noble and Fuhrman (5). in which the Anodisc containing filtered material was submerged in 0.25% SYBR Green I working solution (an optical density of 0.42 at 494 nm when the stock is diluted 1000-fold in sterile deionized water) for 15 min in the dark. Following staining, the filter was removed from the stain, and additional moisture was carefully removed using KimWipes® (Kimberly-Clark, Bedfordview, Gauteng, South Africa). The Anodisc filter was then mounted on a glass slide containing a drop of 50% glycerol, 50% phosphate-buffered saline (PBS; 0.05 M Na2HPO4, 0.85% NaCl, pH 7.5) with 0.1% phenylene-diamine, and covered with a 25-mm square coverslip prior to viewing under oil immersion (1000× magnification) under (UV) blue excitation using a Zeiss Axioskop® 2 plus microscope (Carl Zeiss, Poughkeepsie, NY, USA) as described (5). The heat-treated T4 samples were still detectable by epifluorescent microscopy (Figure 2C) similar to the untreated control (Figure 2A).





We then investigated whether the destabilized capsids were DNase I sensitive. Both the heat-treated and untreated capsids were administered with DNase I as follows: CsCl-purified T4 stock (1 × 106 pfu/mL) in buffer [1 mM Tris-HCl, pH 7.5, 1 mM MgCl2, 5 mM NaCl, 0.01 mg/mL bovine serum albumin (BSA)] was combined with 3 U DNase I (Fermentas, Hanover, MD, USA), incubated for 60 min at 37°C and visualized by epifluorescent microscopy as described. Untreated capsids continued to fluoresce following DNase I treatment (Figure 2B), similar to previous observations with environmental phage samples (10), while heat-treated capsids lost the fluorescent signal following DNase I treatment (Figure 2D), confirming that the heat-treated capsids were DNase I accessible.

In conclusion, we showed that heat-treated T4 capsids retained the gross structural features of the virion but appeared deformed. Furthermore, epifluorescent microscopy revealed that exposure of heat-treated T4 capsids to DNase I abrogates the fluorescent signal that can then be detected in situ without the requirement for extracting DNA from the capsid. Together, these observations suggest a simple method for evaluating capsid structural deformation using epifluorescent microscopy in conjunction with DNase I. The assay should be applicable to any treatment or condition that might alter the capsid's accessibility to DNase I—the hallmark of destabilized capsids. Most dsDNA viruses and phages are DNase I resistant in their native form (11,12,13), and therefore the method should be generally applicable to these DNA-based viruses.

Acknowledgments

The authors are indebted to Rodney Hull for his excellent technical support with the epifluorescent microscope; Carolyn Lalkin for technical support with the electron microscope; Dr. Monde Ntwasa for advice; the National Research Foundation (NRF) for financial assistance; and Tracy McLellan, Denise Lindsay, Digby Warner, and Nathalie Fernandes for critically reviewing the manuscript.

Competing Interests Statement

The authors declare no competing interests.

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