To simulate relevant temperature conditions, publicly available weather data was mined to generate a daily temperature profile for RVF transmission season in Rwanda, May through July, per the 2018 outbreak [1]. A data scraper collected temperature data in half-hour increments from May 18th through July 12th^, 2018, and the temperatures from the same time point each day were averaged to make a single average day. The environmental chamber was then programed to mimic these conditions and a digital thermometer was placed within to confirm the temperature (Supplementary Figure S2). Humidity was maintained at approximately 85% during the experiments roughly matching historical humidity in Rwanda [44].

BUNV was obtained from the World Arbovirus Reference Center at UTMB had been passaged through suckling mouse cells 47 times and Vero cells 6–8 times prior to the experimentation. Stocks were titered via crystal violet plaque assay and determined to be ∼5.0 × 10⁶ pfu/mL. Aedes aegypti mosquito (Rockefeller colony) eggs from were vacuum hatched for 45 min in ddH₂0 and allowed to emerge as in [45].

Viral RNA extraction was performed via MagMax 96 viral isolation as in [43]. qRT-PCR was performed on a Roche Lightcycler 96, using Quantbio Ultratough master mix and previously published primers and probes [43]. Viral RNA quantification was analyzed via comparison to previously crystal violet plaqued stock from above. Standard crystal violet plaque assays were performed [46]. Virus titer was averaged from countable plaques at each dilution [47].

Ae. aegypti reared as above were allowed to mature to adults. Adult mosquitoes were cold anesthetized and separated by sex and placed in to 900 µL BA-1 media with 2 stainless steel BBs in a 2 mL locking Eppendorf tube in pools of 10 and homogenized as previously described [45] and a known titer of virus was added to each tube of titers of 10⁵, 10³, 10¹, and 10⁰ pfu/mL. Five replicates per pools sex were performed at each titer.

Adult Aedes aegypti females, 2 days post emergence (dpe) were offered an infectious blood meal of ratio 2:1 of bovine blood in Alsevers (Hemostat Labs, Dixon, CA, United States) and BUNV infectious supernatant (total infectious titer 1.7 × 10⁶) via the Hemotek feeding apparatus (Discovery Labs, Blackburn, United Kingdom). Engorged females were separated by aspiration and cold anesthetization, then placed in individual 4-ounce containers and provided 10% sucrose solution ad libitum. Each container was supplied with egg paper that was moistened daily and collected when eggs were observed for an additional 18 days. Three females died prior to the end of the experiment and were collected and tested for virus in the legs and bodies as described below. At 18 days post infectious bloodmeal, mosquitoes were processed to determine infection status of bodies, legs, and saliva as in [48].

There is mixed evidence for the role of second bloodmeals in enhancing vector competence [48–51]. Thus, a second cohort of mosquitoes was offered an infectious bloodmeal as above at 3–4 days post emergence. A second uninfected bloodmeal was offered starting at 4 days following the first bloodmeal [50] and repeated attempts were made until all females had taken a second bloodmeal (maximum 9 days following first bloodmeal). The mosquitoes were maintained as described above until processed following egg collection at 22–24 post initial, BUNV infectious bloodmeal.

Ae. aegypti were hatched and reared as described above. Fourth instar larva were collected and rinsed in ddH₂0 and placed into water contaminated with BUNV. Briefly, BUNV supernatant was added to rearing water, and samples were collected at 0–5 days to establish the presence of BUNV RNA [34]. Exposed larva were allowed to mature and were collected as pupa, rinsed, and transferred to a cage for emergence. Pupa were separated based on number of days spent exposed to infected rearing water. Upon emergence, adult mosquitoes were cold anesthetized and separated by sex into pools of up to five based on exposure time. Remaining unemerged individuals were collected at 6 days post contaminated water exposure, rinsed as above, and collated into pools of up to ten for testing by qRT-PCR.

To assess environmental stability of BUNV under environmentally relevant conditions, 110 𝛍L of virus was added to 4.39 mL of two different substrate types–either whole blood or serum for a final concentration of 1 × 105 pfu/mL. The virus-substrate mixture was added to individual units (either falcon tubes or 6-well plates, see Supplementary Methods) and placed into the environmental chamber under Rwanda temperature conditions. Tubes were uncapped and the six-well plate lids lifted to allow air exposure; all units were placed in secondary containment for safety and contamination avoidance. Tubes were sampled at days 0, 1, 7, 21, and 28, while plates were sampled each day until dry. Plates were sampled daily until dry, which occurred between 6 and 8 days. Collected samples were placed into a −80°C freezer until further analysis. Samples were tested starting with the latest timepoint (tubes) or with the day before the plate was dry (Dry Day −1). If a sample was found not to be infectious at a timepoint (I.e., timepoint X), the timepoint immediately preceding (i.e., timepoint X-1) was tested and this pattern continued until we found the latest time of infectivity. Ten replicates of each unit type were conducted per substrate type. Samples were assayed for stability of RNA via qRT-PCR as above and for infectivity via a growth assay on Vero cells as in [43].

For the sensitivity assay, a Kruskal Wallace non-parametric analysis of variance was used to determine if a difference in detection ability existed between sexes of the pools. A linear regression model was used to evaluate the relationship between the initial input into the mosquito pool and the qRT-PCR output. The limit of detection (LOD) was determined using a qPCR LOD calculation R script (at 95% confidence) [52, 53]. The lowest concentration of consistently detectable RNA was determined by running multiple iterations of a standard curve assay tied to a plaque assay. The plaque assay revealed that maximum stock titer was 5.3 × 10⁶ PFU/mL. The standard curve runs determined that the consistent point at which there was below 95% confidence detection was 5.3 × 10⁻¹ PFU/mL. Growth was assessed via non-parametric t-test between day 1 and day 7 where a significantly higher titer at day 7 indicated growth and thus infectiousness. To evaluate stability, RNA recovered at each timepoint was compared via a Kruskal-Wallace test where day was a factor; a post hoc Dunn’s test (Bonferroni correction) test was performed where appropriate. Additionally, comparisons between RNA recovered from each substrate at the same timepoint or day until dry were done via Wilcoxon Ranked Sum tests. Significance was assessed at the α = 0.05 level. Analyses were performed using R Studio (2023.03.1 + 446) and R (4.3.0) and associated packages [54]. Minimum infection rate was determined by calculating the rate of infection as if only one individual per positive pool was infected, as in [55].

The qRT-PCR assay used herein was highly sensitive for BUNV in the context of mosquito pool testing (LOD = 6.6 copies/mL with 95% confidence). There was no significant difference in the amount of viral RNA detected in pools of males versus females (Supplementary Figure S3, p > 0.05), indicating the assay was sensitive in pools of both sexes. There was only one discrepant result where 5/5 male pools versus 4/5 female pools were positive at the lowest titer of 10⁰ pfu/mL. Combining sexes, at titers of 10¹ and above, successful detection of virus was 100% (n = 10). At 10⁰, BUNV was detected in 90% of pools (n = 10, p > 0.05).

We found our colony of Ae. aegypti were not competent for BUNV and thus not able to horizontally or vertically transmit the virus after a single bloodmeal (n = 27). A single mosquito that died at 1 day post exposure tested positive for a midgut infection, but was not included in the analysis as this was more than likely residual RNA from the infectious bloodmeal (midgut titer 1.3 × 10³ pfu/mL). Mosquitoes receiving a second bloodmeal also showed no BUNV midgut infection (n = 19).

Rearing water contaminated with BUNV remained positive for RNA for the 5 days monitored. Initial quantification of RNA copies were approximately at 10^4.5 genome equivalents/mL on days 0–1, while this dropped and remained consistent at 10^2.5–3 for days 2–5. Attempts to quantify BUNV in rearing water via plaque assay were unsuccessful. Despite detection of BUNV RNA, no adult pool tested positive for BUNV. Similarly, a single pool of two pupa that had not emerged by day 6 were also negative for virus (Table 1, see Supplementary Methods). Additionally, larva that had not pupated by day 6 were also pooled and tested. Of the 19 pools, 5 (26%) tested positive for BUNV RNA with low titer (Table 1).

Under simulated Rwandan conditions, in both whole blood and serum, BUNV RNA was detectable at moderate genome copies/mL up to 28 days in tubes (Figure 1A). The difference in average titer of genome copies between whole blood and sera was statistically significant at all time points (p < 0.05). Of interest, both in whole blood and sera, BUNV RNA remained moderately high out to day 28 (6.06 × 10³ and 1.89 × 10⁵, respectively). RNA was stable over all days collected for sera. In whole blood a significant decrease in RNA compared to baseline at day 0 was observed starting on day 21, but no further significant decrease was observed between days 21 and 28 (Figure 1A, p < 0.05). Unsurprisingly, liquid volume in plates evaporated quickly relative to tubes (Supplementary Figures S4–S5) and plate wells were dry between 6 and 8 days post inoculation. The last day of sampling is referred to as Day Dry—1 (Dry-1, see Supplemental Methods). On average, 4.22 × 10⁴ BUNV genome copies/mL was recovered on Dry-1 for whole blood, while the average recovered genome copies in sera was significantly more (p < 0.05) at 1.21 × 10⁵ copies/mL on Dry-1 (Figure 1B). In sera the genome copies were stable as no significant differences were found between the RNA recovered on day 0 and Dry-1 (p > 0.05), while the RNA copies recovered in whole blood increased from day 0 to Dry −1 (p < 0.05). In a subset of plates (n = 5), media was added to dried out wells, collected, and subsequently tested for BUNV RNA (see Supplementary Methods). BUNV RNA was detected in the reconstituted samples, with an average of 6.87 × 10⁴ RNA copies/mL in the reconstituted sera samples, and 6.74 × 10⁴ RNA copies/mL in the reconstituted whole blood samples.

For whole blood in tubes, the last day of infectiousness was day 28 for 80% (8/10) of samples and day 21 for the remining 20%. In tubes containing sera, the last day of infectiousness was day 21 for 50% (5/10) of replicates while the other 50% retained infectiousness out to day 28. Cytopathic effects (CPE) were observed in all infectious samples. Attempts to plaque sera and whole blood at these later time points were unsuccessful due to the consistency of the samples (see Supplementary Methods, Supplementary Figure S4). In 6-well plates, the range of days where infectiousness was retained was 5–7 days post inoculation but was dependent on the remaining volume in the well (Supplementary Figure S3; Supplementary Table S1). Relative to the first day of no volume, 50% (5/10) of the whole blood replicates retained infectiousness at Dry-1, and 100% at Dry-2. For sera however, all replicates retained infectiousness at Dry −1. We were able to plaque plate samples to further investigate the infectious titer of environmentally stable BUNV on the last day of infectiousness. Overall, whole blood had a significantly lower average infectious titer (1.04 × 10³ PFU/mL) compared to sera (2.45 × 10⁴ pfu/mL) (Figure 2, p < 0.05). No CPE was observed in vitro for any of the reconstituted dried samples, indicating a lack of infectivity despite moderate quantitative recovery of ∼10⁴ RNA copies/mL (see Supplementary Table S2).