Conversely, studies examining early infection of mosquitoes by WNV and VEEV demonstrated that only a few (,15) midgut cells are susceptible to arbovirus infection [22,23]. These findings suggest that anatomical barriers, specifically cells of the midgut, may act as genetic bottlenecks by restricting the population of infecting virions thereby diminishing the genetic diversity of the population. Importantly, these observations are not mutually exclusive as several viral genomes may coinfect a single midgut cell. Importantly, population bottlenecks associated with mosquito transmission have not been assessed from a virus genetics perspective. Therefore, we determined whether WNV experiences genetic bottlenecks during the EIP in the vector mosquito Culex pipiens quinquefasciatus. We hypothesized that WNV experiences genetic bottlenecks during the EIP in mosquitoes, and reasoned that sequential reductions in viral genetic diversity would occur as infection progressed throughout the mosquito. To assess this, WNV genetic diversity was quantified in mosquito midguts, hemolymph, and salivary secretions, compartments that represent three well-characterized infection stages (midgut colonization, dissemination, and transmission). Three mosquitoes at three time points (7, 14, and 21 days post infection (dpi)) were sampled. Our data suggest that stochastic reduction of genetic diversity in mosquitoes is at most a minor component of WNV population biology during horizontal transmission. Results Mosquito Infection Rates Mosquito tissues were screened for the presence of WNV RNA by one-step RT-PCR. All freshly fed mosquitoes were positive for WNV RNA representing ‘input’, blood-meal associated virus. The infection rates for midguts at 7, 14, and 21 days post infection (dpi), reflecting viral populations able to overcome the midgut infection barrier, were 88% (21/24), 86% (19/22), and 70% (14/ 20), respectively. The percentage of mosquitoes positive for WNV RNA in the legs, indicating virus dissemination from the midgut and into surrounding hemolymph, was 58% (14/24), 36% (8/22), and 55% (11/20) at 7, 14, and 21 dpi, respectively. In order for mosquitoes to transmit WNV, the virus must be able to overcome the salivary gland infection and escape barriers. The percentage of mosquitoes with WNV in salivary secretions was 25% (6/24), 14% (3/22), and 35% (7/20) at 7, 14, and 21 dpi, respectively. Three mosquitoes per time point with WNV RNA in midgut, hemolymph and saliva were selected for further analysis and WNV genome equivalents quantified (Text S1). Genome equiv- alents were highest in midguts and progressively decreased in the hemolymph and saliva. Further, genome equivalents increased with time post infection (21 dpi.14 dpi.7 dpi) (Figure S1). In addition, three mosquitoes, representing the ‘input’ group, were collected immediately post-engorgment. WNV genome equiva- lents determined for each of these individuals were 3.2, 4.1 and 5.96105 genome equivalents/ml (Figure S1). The bloodmeal contained 66106 pfu/ml, assuming 10–100 genomes per infec- tious particle and an engorgment volume of ,3 ml, engorged mosquitoes would be expected to contain ,1.46105 or 6 genome equivalents. The concentrations for the three individuals in the ‘input’ group are in agreement with these calculations and thus faithfully represent the population of the bloodmeal as a whole. Some arboviruses may enter mosquito hemolymph directly, bypassing midgut infection via a ‘leaky midgut’ [24,25]. In order to determine whether this occurred in the WNV-Cx. quinquefasciatus system, hemolymph was removed from mosquitoes at 1, 3, 24, and 48 hpi as well as 8 and 16 dpi and tested for WNV by plaque assay WNV Diversity is Maintained in Mosquitoes (Text S1). Hemolymph collected at 8 and 16 dpi commonly held high titers of WNV. In contrast, hemolymph collected at early timepoints after feeding almost never contained infectious WNV (Figure S2). WNV Genetic Diversity The percent nucleotide diversity and proportion of unique viral variants were used as indicators of viral genetic diversity in each of the samples. The percent nucleotide diversity was determined by calculating the total number of nucleotide changes for all clones within a given sample divided by the total number of nucleotides sequenced per sample. The data was grouped either by days post infection (Figure 1 A, B, and C) or by tissue type (Figure 1 D, E, and F). Analysis of the data set by days post infection revealed that there was no significant difference in the percent nucleotide diversity among the viral populations sequenced at 7 and 14 dpi between ‘input’, midgut, legs or saliva (p = 0.2739 and p = 0.2662, respectively) (Figures 1 A & B). Interestingly, genetic diversity seemed to decrease with time post infection as there was a significant reduction in diversity from the ‘input’ to the three tissue types at 21 dpi (ANOVA p = 0.0015; Tukey’s HSD post test, ‘input’ vs midgut q = 7.262 p,0.05, ‘input’ vs legs q = 8.493 p,0.05, and ‘input’ vs saliva q = 5.293 p,0.05), but no difference between tissue types (Figure 1C). Analyzing the data by tissue type revealed that there was a significant reduction in diversity from the ‘input’ to midguts at 14 and 21 dpi (ANOVA p = 0.0125, Tukey’s HSD post test, ‘input’ vs 14 dpi q = 5.404 p,0.05 and ‘input’ vs 21 dpi q = 5.694 p,0.05), but no significant difference between midguts at 7 dpi and ‘input’ (Figure 1D). There was no statistical differences between any of the leg or saliva samples at 7, 14 or 21 dpi (legs p = 0.0996, saliva p = 0.3563) (Figure 1E & 1F). The second indicator of genetic diversity used in these studies was the proportion of unique viral variants. This was determined by calculating the number of unique clones per sample and dividing by the total number of clones sequenced per sample. Again the data was grouped either by days post infection (Figure 2 A, B, & C) or tissue type (Figure 2 D, E, & F). At 7 dpi, the midguts and saliva were significantly lower than the ‘input’ (ANOVA p = 0.0052, Tukey’s HSD post test, ‘input’ vs midguts q = 7.038 p,0.05, ‘input’ vs saliva q = 5.841 p,0.05), but the tissues were not significantly different from one another (Figure 2A). Interest- ingly, by 14 dpi the proportion of unique viral variants between the tissues and ‘input’ was not significant (ANOVA p = 0.0517), but at 21 dpi each of three tissue types were significantly lower than the ‘input’ (ANOVA p = 0.0021, Tukey’s HSD post test, ‘input’ vs midguts q = 6.898 p,0.05, ‘input’ vs legs q = 8.028 p,0.05, and ‘input’ vs saliva q = 5.306 p,0.05) (Figure 2B & 2C). Analysis by tissue type revealed that midguts from all three time points were significantly lower than the ‘input’, but not different between time points (ANOVA p = 0.0014, Tukey’s HSD post test, ‘input’ vs 7 dpi q = 7.298 p,0.05, ‘input’ vs 14 dpi q = 7.838 p,0.05, and ‘input’ vs 21 dpi q = 7.576 p,0.05) (Figure 2D). Like the midguts, the legs at 14 and 21 dpi contained significantly less diversity than the ‘input’ (ANOVA p = 0.0018, Tukey’s HSD post test, ‘input’ vs 14 dpi q = 6.597 p,0.05 and ‘input’ vs 21 dpi q = 8.458 p,0.05), but was not different from the 7 dpi time point. Further, there was no difference between the time points (Figure 2E). Finally, comparison of the saliva samples at each of the time points revealed no significant difference between the three time points and the ‘input’ or between time points (ANOVA p = 0.1431) (Figure 2F). Because our frequency and location analysis of viral variants revealed that numerous variants were found in both the ‘input’ and saliva, but not the midgut or legs, we performed a correlation PLoS ONE | www.plosone.org 2 September 2011 | Volume 6 | Issue 9 | e24466