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\begin{document}

\title{Human vaccine candidate (DDVax) development against Rift Valley Fever:
dissemination safety studies in mosquitoes}



\author[1]{Corey L. Campbell}%
\author[1]{Trey K. Snell}%
\author[1]{Susi Bennett}%
\author[2]{John Wyckoff}%
\author[1]{Emma K. Harris}%
\author[1]{Daniel A. Hartman}%
\author[1]{Elena Lian}%
\author[3]{Brian Bird}%
\author[1]{Mark D. Stenglein}%
\author[1]{Richard Bowen}%
\author[1]{Rebekah Kading}%
\author[2]{* Darragh}%
\affil[1]{Colorado State University}%
\affil[2]{BioMARC Infectious Diseases Research Center CSU}%
\affil[3]{University of California Davis School of Veterinary Medicine}%


\vspace{-1em}



  
  \date{May 25, 2024}


\begingroup
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\selectlanguage{english}
\begin{abstract}
Rift Valley fever virus (RVFV) is a mosquito-borne pathogen with
significant human and veterinary health consequences that periodically
emerges in epizootics. RVF causes fetal loss and death in ruminants and
in humans can lead to liver and renal disease, delayed-onset
encephalitis, retinitis, and in some cases severe hemorrhagic fever. A
live attenuated vaccine candidate (DDVax), was developed by the deletion
of the virulence factors NSs and NSm from a clinical isolate, ZH501, and
has proven safe and immunogenic in rodents, pregnant sheep and non-human
primates. Deletion of NSm also severely restricted mosquito midgut
infection and inhibited vector-borne transmission. To demonstrate
environmental safety, this study investigated the replication,
dissemination and transmission efficiency of DDVax in mosquitoes
following oral exposure compared to RVFV strains MP-12 and ZH501.
Infection and dissemination profiles were also measured in mosquitoes 7
days after feeding on goats inoculated with DDvax or MP-12. Hypothesis:
DDVax should infect mosquitoes at significantly lower rates than other
RVF strains and, due to lack of NSm, be transmission
incompetent.~Exposure of \emph{Ae. aegypti} and \emph{Cx. tarsalis} to
6-8 log \textsubscript{10} plaque forming units (PFU)/mL DDVax by
artificial bloodmeal resulted in significantly reduced DDVax infection
rates in mosquito bodies compared to controls. Plaque assays indicated
negligible transmission of infectious DDVax in \emph{Cx. tarsalis}
saliva (1/140 sampled) and none in \emph{Ae aegypti} saliva (0/120).
Serum from goats inoculated with DDVax or MP-12 did not harbor
detectable infectious virus by plaque assay at 1, 2, or 3
days-post-inoculation; infectious virus was, however, recovered from
mosquito bodies that fed on goats vaccinated with MP-12 (13.8\% and
4.6\%, respectively), but strikingly, DDvax positive mosquito bodies
were greatly reduced (4\%, and 0\%, respectively). Furthermore, DDVax
did not disseminate to legs/wings in any of the goat-fed mosquitoes.~
Collectively, these results are consistent with a beneficial
environmental safety profile \emph{.}%
\end{abstract}%



\sloppy


\textbf{Title-} Human vaccine candidate (DDVax) development against Rift
Valley Fever:~dissemination safety studies in mosquitoes

\textbf{Authors-} Corey L. Campbell\selectlanguage{ngerman}\textsuperscript{1§}, Trey K.
Snell\textsuperscript{1}, Susi Bennett\textsuperscript{1}, John
Wyckoff\textsuperscript{2},~Darragh~Heaslip\textsuperscript{2}, Emma K.
Harris\textsuperscript{1}, Daniel A. Hartman\textsuperscript{1}, Elena
Lian\textsuperscript{1}, Brian H. Bird\textsuperscript{3}, Mark D.
Stenglein\textsuperscript{1}, Richard A. Bowen\textsuperscript{1},
Rebekah C. Kading\textsuperscript{1}

\textsuperscript{1.}Center for Vector-borne Infectious Diseases,
Department of Microbiology, Immunology, and Pathology, Colorado State
University. \textsuperscript{2.}BioMARC, Infectious Diseases Research
Center, CSU.

\textsuperscript{3} One Health Institute, School of Veterinary Medicine,
University of California, Davis, CA, USA.

\textsuperscript{§}Corresponding author: Corey L. Campbell, Colorado
State University, Campus Delivery 1685, Fort Collins, CO 80523, USA.

Keywords: vaccine, vector competence, arbovirus, vector-borne disease,
Aedes, Culex

email- C. L. Campbell, corey.campbell@colostate.edu

email- T. K. Snell, tksnell78@gmail.com

email- S. Bennett, susi.bennett@colostate.edu

email- J. Wyckoff, john.wyckoff@colostate.edu

email- D.~Heaslip, darragh.heaslip@colostate.edu

email- E. K. Harris, emkate.harris@colostate.edu

email- D. A. Hartman, hartman.daniel.a@gmail.com

email- E. Lian, elian@rams.colostate.edu

email- B. H. Bird, bhbird@ucdavis.edu

email- M. D. Stenglein, mark.stenglein@colostate.edu

email- R.A. Bowen, rbowen@rams.colostate.edu

email- R. C. Kading,\emph{rebekah.kading@colostate.edu}



\section*{Summary}

{\label{summary}}

Rift Valley fever virus (RVFV) is a mosquito-borne pathogen with
significant human and veterinary health consequences that periodically
emerges in epizootics. RVF causes fetal loss and death in ruminants and
in humans can lead to liver and renal disease, delayed-onset
encephalitis, retinitis, and in some cases severe hemorrhagic fever. A
live attenuated vaccine candidate (DDVax), was developed by the deletion
of the virulence factors NSs and NSm from a clinical isolate, ZH501, and
has proven safe and immunogenic in rodents, pregnant sheep and non-human
primates. Deletion of NSm also severely restricted mosquito midgut
infection and inhibited vector-borne transmission. To demonstrate
environmental safety, this study investigated the replication,
dissemination and transmission efficiency of DDVax in mosquitoes
following oral exposure compared to RVFV strains MP-12 and ZH501.
Infection and dissemination profiles were also measured in mosquitoes 7
days after feeding on goats inoculated with DDvax or MP-12. Hypothesis:
DDVax should infect mosquitoes at significantly lower rates than other
RVF strains and, due to lack of NSm, be transmission
incompetent.~Exposure of \emph{Ae. aegypti} and \emph{Cx. tarsalis} to
6-8 log\textsubscript{10} plaque forming units (PFU)/mL DDVax by
artificial bloodmeal resulted in significantly reduced DDVax infection
rates in mosquito bodies compared to controls. Plaque assays indicated
negligible transmission of infectious DDVax in \emph{Cx. tarsalis}saliva
(1/140 sampled) and none in \emph{Ae aegypti} saliva (0/120). Serum from
goats inoculated with DDVax or MP-12 did not harbor detectable
infectious virus by plaque assay at 1, 2, or 3 days-post-inoculation;
infectious virus was, however, recovered from mosquito bodies that fed
on goats vaccinated with MP-12 (13.8\% and 4.6\%, respectively), but
strikingly, DDvax positive mosquito bodies were greatly reduced (4\%,
and 0\%, respectively). Furthermore, DDVax did not disseminate to
legs/wings in any of the goat-fed mosquitoes.~ Collectively, these
results are consistent with a beneficial environmental safety
profile\emph{.}

\section*{1 Introduction}

{\label{introduction}}

Rift Valley fever virus (RVFV) (family \emph{Phenuiviridae} ,
genus\emph{Phlebovirus} ) is a mosquito-borne virus that causes periodic
epizootic outbreaks across Africa and the Arabian peninsula(Al-Afaleq \&
Hussein, 2011; Nguku et al., 2010). In ruminants, primarily sheep,
goats, camels and camelids, RVF is often characterized by sudden
epizootics marked by near universal fetal death at all stages of
gestation, congenital malformations(Coetzer, 1982) and significant adult
animal deaths often due to acute virus induced hepatic and renal
pathology(Wichgers Schreur et al., 2021). Though most human cases are
typically self-limiting with mild to moderate symptoms (McElroy, Harmon,
Flietstra, Nichol, \& Spiropoulou, 2018), cases of delayed onset
encephalitis, kidney and/or eye damage, severe anemia, hemorrhagic fever
and miscarriage can occur(Baudin et al., 2016; Coetzer, 1982; Madani et
al., 2003; Oymans, Wichgers Schreur, van Keulen, Kant, \& Kortekaas,
2020).

Over 40 species of mosquitoes have been demonstrated as competent
vectors for RVFV(reviewed in(Lumley et al., 2017)), some of which range
on multiple continents. Following periods of heavy rainfall, which
stimulate rapid increases in vector mosquito populations, RVFV
re-emerges periodically in explosive epizootics(Al-Afaleq \& Hussein,
2011; Nguku et al., 2010). In the absence of humans and livestock, RVFV
cycles between mosquitoes and wild ruminants(Britch et al., 2013; Clark,
Warimwe, Di Nardo, Lyons, \& Gubbins, 2018). Between epizootics, there
is also support for low level maintenance of RVFV in livestock in
inter-epidemic periods(Lichoti et al., 2014).

Due to these health implications and the potential to cause a public
health emergency, in 2018 the World Health Organization listed RVFV as a
research and development blueprint priority pathogen(Mehand,
Al-Shorbaji, Millett, \& Murgue, 2018). Availability of a safe and
effective human vaccine against RVFV is essential to protect the health
of people in endemic regions and a preparatory measure for the
anticipated cross-border spread and establishment of RVFV into new
geographic areas. A number of vaccine candidates have been developed for
RVFV, including formalin inactivated (Pittman et al., 1999; Randall,
Gibbs, Aulisio, Binn, \& Harrison, 1962) and live attenuated
strains(Ikegami et al., 2015; Smithburn, 1949). However residual
teratogenic effects in animals or the need for boosters to maintain
protective immunity(Bird, Ksiazek, Nichol, \& Maclachlan, 2009; Botros
et al., 2006) present challenges for further development of these
earlier candidates. To date, there is currently no commercially
available and fully FDA-approved RVFV human vaccine.

To meet this critical health need, a human vaccine candidate (DDVax), a
double deletion construct of the parental wild-type strain ZH501 was
generated using a reverse genetics approach wherein both the NSs
(non-structural, S segment) and NSm (non-structural, M segment)
virulence genes were removed(Bird et al., 2008). NSs is expressed in an
ambisense fashion from the viral S segment(Ikegami et al., 2009) and is
a multi-functional protein that antagonizes host cell interferon
responses(Le May et al., 2008). The viral M segment encodes 2 major
glycoproteins and multiple open reading frames in the NSm coding
regions, which is required for efficient dissemination in
mosquitoes(Crabtree et al., 2012). Neither NSs nor NSm are required for
viral replication in interferon-deficient cell culture, and the
attenuated DDVax vaccine candidate has proven to be safe and immunogenic
in a variety of animal species with the added benefit of inhibited
replication and transmission in mosquitoes(Bird et al., 2008; Bird et
al., 2011; Crabtree et al., 2012; Kading et al., 2014).

The objective of this study was to confirm that the newly rescued
version of DDVax produced for development under Good Manufacturing
Practices (GMP) behaved as previously described and exhibited a highly
favorable environmental safety profile by not being transmitted by
potential mosquito vectors. Here, we describe characterization of RVFV
DDVax in mosquitoes using both \emph{in vitro} and \emph{in
vivo}approaches. These vector assessments were divided into two
experimental phases: 1) mosquito oral challenges via artificial feeding
and 2) mosquito feeding on DDVax inoculated goats. Features of vector
competence were measured in two competent mosquito species, \emph{Culex
tarsalis} Coquillett and \emph{Aedes aegypti} L., to determine
infection, dissemination and transmission potential, using reverse
transcriptase- quantitative PCR (RT-qPCR) and infectious virus plaque
assay. Vertebrate-to-vector transmission from DDVax-inoculated goats to
mosquitoes was also measured. Collectively, these experiments provided
important comparison of vector competence of mosquitoes exposed to
DDVax(Bird et al., 2008), ZH501, the parental wild-type virus and MP-12,
an existing vaccine virus strain(Turell \& Rossi, 1991).

\section*{2 Methods}

{\label{methods}}

\subsection*{2.1 DDVax Production Process
Summary}

{\label{ddvax-production-process-summary}}

\subsubsection*{2.1.1 Uninfected Cell
Culture}

{\label{uninfected-cell-culture}}

Serum-Free Vero cells (2 x
10\textsuperscript{4}cells/cm\textsuperscript{2}) were grown in OptiPro
Serum-Free Media (SFM) (Lifetech, A31343) with Glutamax (Gibco,
35050-061) at 37°C and 5\% CO\textsubscript{2}. Cells were expanded into
3 x 10-Layer CellStacks (6360 cm\textsuperscript{2}) (Corning, 3271) and
1 x 1-Layer CellStacks (636 cm\textsuperscript{2}) (Corning, 3268). For
each passage, cells were seeded at either 2.0 x
10\textsuperscript{4}cells/cm\textsuperscript{2} for 48 ± 8-hours or 1.5
x 10\textsuperscript{4} cell/cm\textsuperscript{2} for 72 ± 8-hours.
Cell harvest was performed using TrypLE Select (Life Tech, 12563092).
Cells were centrifuged at 500 x g for 5 minutes at 18°C and resuspended
in OptiPro SFM. Cell enumeration was performed using a Vi-Cell Cell
Viability Analyzer.

\subsubsection*{2.1.2 Generation of DDVax Pilot
Material}

{\label{generation-of-ddvax-pilot-material}}

Upon achieving cell confluency within the 3 x 10-Layer or 1 x 1-Layer
CellStacks, Vero cells were infected with the DDVax Research Virus Stock
(RVS), Lot \# N16-5-20-RV at a multiplicity of infection (MOI) of 0.0005
PFU/cell. Infection volume used for each 10-Layer CellStack was 1300ml
and the infection volume for the 1-Layer CellStack was 130ml. Infected
cultures were then incubated at 37°C and 5\% CO\textsubscript{2} for
72-hours. Following the 72-hour infection incubation, flasks were
examined for cytopathic effects (CPE). DDVax virus was then harvested by
pumping the supernatant from each CellStack into a 5L Flexboy bag. The
harvested pool underwent Benzonase treatment to digest Host Cell DNA
(HCD). A 500mM MgCl\textsubscript{2} solution was added to the Flexboy
bag to achieve a final concentration of 1.5mM MgCl\textsubscript{2} for
Benzonase treatment. Benzonase (EMD Millipore, 101679) was added at a
concentration of 50,000 U/L. The Flexboy bag was thoroughly rocked to
mix and then incubated for 60 minutes at 37°C and 5\%
CO\textsubscript{2}, with rocking at 10-minute intervals. The
Benzonase-treated pool was divided into 500ml conical tubes and
centrifuged at 3000 x g for 15 minutes at 18°C to remove large cell
debris. Supernatant containing the DDVax, now the clarified pool, was
pumped out of the centrifuge bottles into a new 5L Flexboy bag. Virus
was then concentrated by a factor of 6 using ultrafiltration (UF) using
the KMPi TFF system (Repligen) over a 500kD Hollow Fiber Membrane
(Repligen, S02-E500-05-N, 500kD, 20cm length, 0.5mm fiber diameter).
Concentration was performed at a target shear rate of 3000
s\textsuperscript{-1} and TMP setpoint of 5 psi. The concentrated UF
pool was diafiltered (DF) into a buffer containing 0.2M NaCl, 10mM
Sodium Phosphate, 4\% Sucrose, 5mM Glutamax, pH 7.4 ± 0.1. Buffer
exchange was performed for 10 diavolumes. Diafiltration was performed at
a target shear rate of 3000 s\textsuperscript{-1}

\subsection*{2.2 DDVax sequencing and
analysis}

{\label{ddvax-sequencing-and-analysis}}

Illumina shotgun sequencing libraries were prepared from total RNA using
the Kapa RNA HyperPrep kit following the manufacturer's protocol. Dual
indexed libraries were sequenced on an Illumina NextSeq 500 sequencer to
generate single-end 150 nt reads.

We used two complementary approaches to detect and quantify viral
variants. First, we used the lofreq tool to identify single nucleotide
variants and short insertions and deletions (Wilm et al., 2012). Second,
we used DI-tector to identify structural variants including longer
deletions and insertions and copy back defective viral genomes
(DVGs)(Beauclair et al., 2018; Vignuzzi \& Lopez, 2019). These tools
were run as part of a reproducible Nextflow pipeline, available at
https://github.com/stenglein-lab/viral\_variant\_caller/releases/tag/DDVax\_paper\_release.
Software dependencies and reference sequences (DDVax) are captured in
this version-controlled release and in the conda environment contained
therein.

To quantify variants, adapter-derived and low-quality bases were trimmed
using Cutadapt(Martin, 2011). Host cell-derived reads were removed using
bowtie2 to align reads to the \emph{Chlorocebus sabeus} genome,
accession GCF\_000409795.2(Langmead \& Salzberg, 2012). Host- and
quality-filtered reads were aligned to the S, M, and L segment
RVFV/DDVax reference sequences using the BWA aligner(Langmead \&
Salzberg, 2012; Li \& Durbin, 2009). The reference sequences consisted
of the RVFV-derived portions of the DDVax plasmid sequences. To improve
accuracy of structural variant (indel) calls, base quality scores were
recalibrated using GATK (McKenna et al., 2010). Single nucleotide
variants (SNVs) and structural variants were called using LoFreq(Wilm et
al., 2012). The minimum depth of coverage to call a variant was set at
40x coverage. SnpEff and SnpSift were used to predict the functional
impact of variants(Cingolani, Patel, et al., 2012; Cingolani, Platts, et
al., 2012). Variant calling distinguished between variants that were not
detected despite sufficient data and positions that lacked sufficient
data to call variants. Defective viral genomes were identified using the
DI-tector tool(Beauclair et al., 2018). Outputs of these analyses were
tabulated, processed, and visualized in R. Variants with frequencies \selectlanguage{english}[?]
3\% were reported(Grubaugh et al., 2019).

\subsection*{2.3 Virus strains}

{\label{virus-strains}}

Stocks of DDVax were produced as described above. The passage history of
MP-12 strain is unknown, but DQ375404.1 (L segment), DQ380208.1 (M
segment) and DQ380154.1 (S segment) sequences were confirmed by Sanger
sequencing. ZH501 strain virus was obtained from R. Bowen. V1 (Vero)
passage stock was passaged twice in Vero cells to obtain V3 stocks used
for this study.

\textbf{2.4 Insect cell culture virus growth curves}

The insect cell lines used for this study were Ct cells, derived
from\emph{Cx. tarsalis} embryos (Centers for Disease Control and
Prevention)(Chao \& Ball, 1976), Aag2 \emph{Ae. aegypti} high passage
cells, also derived from embryos(Chao \& Ball, 1976), and ATC10 (CCL-125
(ATCC)), a larval-derived cell line(Singh, 1971). Virus strains
RVFV-ZH501, the parental wild-type to DDVax(Bird et al., 2008),
MP-12(Turell \& Rossi, 1991) and DDVax were subjected to growth curves
in mosquito cell culture (ATC-10, Aag2, CT) using Schneider's media
(10\% FBS (or 20\% FBS for ATC-10s), 1\% non-essential amino acids, 1\%
L-glutamine, 1\% penicillin/streptomycin). An MOI of 0.01 was used for
all infections. Aliquots were removed at daily timepoints for 1-6 days
post-infection (dpi). At each timepoint, 400 \selectlanguage{greek}µ\selectlanguage{english}l cell culture supernatant
was removed, and media was replaced into the T-flask. Aliquots were
supplemented with 20\% FBS as a cryoprotectant and stored at -80\selectlanguage{ngerman}°C until
titrations were performed.

\textbf{2.5 Mosquitoes}

The Poza Rica \emph{Ae. aegypti} strain was colonized in 2012 from
collections in the state of Veracruz, Mexico(Vera-Maloof,
Saavedra-Rodriguez, Elizondo-Quiroga, Lozano-Fuentes, \& Black IV,
2015). The \emph{Cx. tarsalis} Kern National Wildlife Refuge (KNWR)
colony(Oviedo, Romoser, James, Mahmood, \& Reisen, 2011), established in
1952, was obtained from the Centers for Disease Control and Prevention
(Fort Collins, CO). Mosquito colonies were maintained at
24-26\textsuperscript{o}C (\emph{Culex} ) or 28\textsuperscript{o}C
(\emph{Aedes} ) at 12:12 light:dark cycle; adults were fed water and
sucrose \emph{ad libitum} . Larvae were reared on ground TetraMin fish
food (http://www.tetra-fish.com/).

\textbf{2.6 Vector competence}

Adult mosquitoes (4-10 days old) were provided an oral, artificial meal
containing freshly grown RVFV. To approximate titers of 7
log\textsubscript{10} PFU/ml, virus was harvested 3 days after infection
of Vero cells at an MOI of 0.01. Viral supernatant was mixed 1:1 in
defibrinated calf blood, with the addition of 1mM ATP, and 0.075\%
sodium bicarbonate. Mosquitoes were fed for 1 to 1 ½ hours using either
a water-jacketed feeder (https://lillieglassblowers.com) for DDVax and
MP-12 or a hemotek (http://hemotek.co.uk/), in the case of ZH501. All
ZH501 feedings and mosquito incubation steps were performed in the
animal biosafety level 3 conditions. All other \emph{in vitro}mosquito
oral exposures were conducted in standard biosafety level 3 containment.
Fully-engorged mosquitoes were separated into cartons and provided
sucrose and water \emph{ad libitum} . Mosquitoes were held for 14 days
at \textasciitilde{}80\% humidity and 28\textsuperscript{o}C. Infectious
blood meal titers were determined through back titration of the
infecting blood meals.

At 14 days post-challenge, mosquitoes were anesthetized at
4\textsuperscript{o}C, then held on ice during processing. Tissue
samples were dissected, then placed in separate tubes of 250 \selectlanguage{greek}µ\selectlanguage{english}l mosquito
diluent (DMEM, 20\% heat-inactivated FBS, 50 \selectlanguage{greek}µ\selectlanguage{english}g/ml Pen-Strep, 50 \selectlanguage{greek}µ\selectlanguage{english}g/ml
gentamicin, and 2.5 \selectlanguage{greek}µ\selectlanguage{english}g/ml amphotericin B), as follows: Legs and wings
were removed for determination of viral dissemination. Saliva was
collected for determination of transmission potential. The mosquito
proboscis was placed in a capillary tube containing type B immersion oil
(Bioworld, SKU- 21750002) and allowed to salivate for 30-60 minutes. At
that time, the capillary tube was removed and placed in a tube
containing mosquito diluent (1x PBS supplemented

with 20\% FBS (heat-inactivated), 50 \selectlanguage{greek}μ\selectlanguage{english}g/ml Penicillin/Streptomycin, 50
\selectlanguage{greek}μ/\selectlanguage{english}ml Gentamycin,

2.5\selectlanguage{greek}μ\selectlanguage{english}g/ml Fungizone) and centrifuged at 14,000 x g for 3 minutes. Lastly,
each remaining body was also placed in a separate tube, for measurement
of infection status. Samples were homogenized on a Qiagen Tissuelyzer
(Qiagen) at 30 beats per second frequency for 30 seconds, then pelleted
at 14,000 x g in a microfuge at 4\textsuperscript{o}C for 3 minutes.
Samples were stored in -80\textsuperscript{o}C.

\textbf{2.7 RNA extractions}

RNA extractions of individual 50\selectlanguage{greek}µ\selectlanguage{english}l sample aliquots were performed using
the Applied Biosystems MagMax-96 Viral RNA extraction kit (AMB1836-5,
Thermofisher) following the manufacturer's protocol for manual
extraction methods (MAN0017826). Linear polyacrylamide was used as a
carrier in place of carrier RNA. Reactions were eluted into 50\selectlanguage{greek}µ\selectlanguage{english}l elution
buffer and stored in 96-well plates at -80\textsuperscript{o}C.

\textbf{2.8 Reverse transcription quantitative PCR (RT-qPCR)}

RT-qPCR was performed in duplicate using 5 \selectlanguage{greek}µ\selectlanguage{english}l sample or RNA standards
and run on a QuantStudio 2.0 qPCR platform (Applied Biosystems).
Calculated virus amounts were adjusted to account for RNA copy number
per tissue. The following primers were used to quantitate RVFV RNA in
all samples: RVFL-2912fwdgg TGAAAATTCCTGAGACACATGG, RVFL-2971revAC
ACTTCCTTGCATCATCTGATG, RVFL-2950-Probe
(FAM)-CAATGTAAGGGGCCTGTGTGGACTTGTG-(BHQ1)(Bird, Bawiec, Ksiazek,
Shoemaker, \& Nichol, 2007). TaqMan Fast Virus 1-Step Master Mix
(Applied Biosystems) was used with final primer concentrations of 500nM
and a probe concentration of 100nM. Samples and standards were loaded
into 96-well plates and run using fast cycling mode on an AB QuantStudio
machine, using the manufacturer's recommended settings. The cycling
conditions were as follows: 50\textsuperscript{o}C 5 min (1 cyc),
95\textsuperscript{o}C 20 sec (1 cyc), (95\textsuperscript{o}C 3 sec,
60\textsuperscript{o}C 30 sec (40 cyc)).

RNA copy number standards were developed by amplifying a portion of the
L segment from 20 ng plasmid bearing the full length gene(Bird et al.,
2008). The RVFL2173\_T7\_F amplification forward primer
(TAATACGACTCACTATAGGGCAGGTGAGCCCTTCATTCT) contained a T7 promoter;
RVFL3542\_R was the reverse primer (GAGGGGTAAATGGCAAGGTACA). 100 ng
input of PCR product was used \emph{in vitro} transcription reactions
that incubated for 5 hours at 37\textsuperscript{o}C using the
manufacturer's recommendations. Transcription products were stored in 5
\selectlanguage{greek}µ\selectlanguage{english}l aliquots at -80\selectlanguage{ngerman}°C; they were quantitated using a Qubit fluorometer
(ThermoFisher) using the manufacturer's recommendations. For RT-qPCR,
fresh aliquots of \emph{in vitro} transcription reactions were serially
diluted in 10-fold increments to generate standard curves to relate copy
number to raw cycle threshold (Ct value). One standards plate was run
for all samples screened on a given day. A representative standard curve
was y= -3.3111x + 36.655 R\textsuperscript{2}= 0.9976, where y= Ct value
and x= log\textsubscript{10} RNA copy number.

\textbf{2.9 DDVax dose response experiment}

A dose response experiment was performed as a follow up to the mosquito
vector competence challenges, which were administered with only a single
high titer of over 8.0 log\textsubscript{10} PFU/ml. The purpose of this
experiment was to test the hypothesis that \emph{Cx. tarsalis} DDVax
infection rates vary as a function of virus titer in the artificial
blood meal. \emph{Cx tarsalis} were exposed to oral bloodmeals at 6.2,
4.5, or 3.5 log\textsubscript{10} PFU/ml and held for 14 days at
28\textsuperscript{o}C, rH 80\%. At 14 days-post-feeding legs/wings,
saliva and bodies were harvested into mosquito diluent as above in
individual tubes and stored at -80°C. Sample processing was performed as
described above.

\textbf{2.10 Goat virus inoculations and mosquito challenge}

Mature female, non-pregnant dairy goats of multiple breeds were acquired
from a commercial dairy and housed in an Animal Bio-Safety Level 3
facility for the duration of the experiment. Goats were inoculated with
5.6 log\textsubscript{10} PFU freshly grown MP-12 or 6.6
log\textsubscript{10} PFU DDVax, as determined by plaque assay. Blood
was drawn from goat jugular vein at days 1, 2, and 3 post-inoculation
into gel serum separator tubes (Becton Dickson, https://www.bd.com/);
serum was collected by spinning at 1200 \emph{x g} for 10 minutes. Serum
was aliquoted and stored at -80\textsuperscript{o}C. Serum samples were
titered by plaque assay, and RNA was extracted for detection and
quantification of viral RNA.

For mosquito feeding, goats were manually restrained, and mosquitoes
were held in cartons with mesh bottoms against patches of clipped fur
and held for about 30 minutes to allow feeding on days 1 and 2
post-inoculation (Figure S1). Since \emph{Cx. tarsalis} mosquitoes did
not feed well on goats, on day 3 post- inoculated, mosquitoes were
exposed in the laboratory to freshly-collected goat blood (collected
into EDTA tubes (Becton Dickson,\emph{https://www.bd.com/)}) using a
water jacketed feeding apparatus heated to 37\textsuperscript{o}C.
Engorged mosquitoes were held for 7 days at 28\textsuperscript{o}C, rH
80\%. At 7 days-post-feeding, bodies and legs/wings were placed in
individual tubes containing mosquito diluent (see above). Samples were
homogenized on a Qiagen Tissuelyzer (Qiagen) at 30 beats per second
frequency for 30 seconds, then pelleted at 14,000 \emph{x g} in a
microfuge at 4\textsuperscript{o}C for 3 minutes. Tubes were stored in
-80\textsuperscript{o}C. Infectious virus (CPE+/-) was measured by
plaque assay using 100 \selectlanguage{greek}µ\selectlanguage{english}l undiluted sample in duplicate to determine the
frequency of mosquito bodies bearing infectious DDVax virus or MP12 RVFV
(control). For those with RVFV-positive bodies, legs/wings were also
titrated by plaque assay to determine the frequency of mosquitoes with
disseminated infectious virus.

\textbf{2.11 Virus titrations}

Vero cells were grown to \emph{\textgreater{}} 95\% confluency in
Dulbecco's modified eagle media DMEM (5\% fetal bovine serum (Atlas
Biologicals), 1\% sodium bicarbonate, 1\% non-essential amino acids, no
phenol red) in 6 or 12-well plates. Ten-fold serial dilutions of virus
stocks and blood meal aliquots in media were performed. Mosquito samples
were used undiluted. \emph{In vitro} challenged mosquito samples had
already undergone one freeze-thaw cycle prior to infectious virus
detection. For each dilution or sample, one hundred microliters of
sample was added to wells, then incubated with rocking for 1 hour,
followed by an overlay (0.4\% agarose (Lonza Rockland) in DMEM). At 2
days post-infection, overlays (0.33\% neutral red (Sigma N2889), 2\%
agarose in supplemented DMEM) were applied. Plates were read after 24
hours. Ambiguous plaques were more closely examined under an inverted
microscope at 40X magnification to better confirm CPE.

\textbf{2.12 Data Analysis}

P was determined by calculating the proportion of viral RNA-positive
mosquito bodies for the combined total number of mosquito RNA samples.
Dissemination was determined by calculating the proportion of legs/wings
RNA samples with detectable RVFV RNA against the total number of
mosquitoes exposed. Transmission was determined by calculating the
proportion of saliva RNA samples that were RVFV-RNA positive against the
total number of mosquitoes exposed. Percent of saliva expectorants
containing infectious virus were also calculated by determining the
proportion of saliva samples producing detectable CPE by plaque assay
among the total number of individuals tested. The percentage of
RVFV-infected mosquitoes after feeding on inoculated goats were
determined by calculating plaque positive mosquito bodies per total
number of mosquitoes assayed. RVFV growth curve titers were analyzed by
calculating the highest dilution containing countable plaques and
multiplying that by the dilution factor to obtain
log\textsubscript{10}PFU/ml.

All graphing and statistical tests were performed in Prism Graphpad
(version 8, https://www.graphpad.com/). \selectlanguage{greek}χ\selectlanguage{english}\textsuperscript{2} contingency
tests were used to calculate dissemination and transmission rates. Two
way ANOVA (analysis of variance) with Geisser-Greenhouse correction was
used to determine differences in viral growth kinetics. One way ANOVA
was used to determine differences in bloodmeal titers.

\section*{3 Results}

{\label{results}}

\textbf{3.1 DDVax variant analysis}

We used sequencing to track the genetic stability of DDVax over 5
passages in Vero cell culture (P1 through P5, MOI 0.0005). The P5
preparation was used for goat inoculations. Total RNA from virus
preparations (supernatant: P1-P4, or filtered supernatant: P5) was
converted into shotgun Illumina libraries and sequenced on an Illumina
NextSeq 500 instrument to produce a median of
1.2x10\textsuperscript{7}single end 150 nucleotide (nt) reads per
sample. After removing low quality and adapter-derived bases, a median
of 1.1x10\textsuperscript{7}reads (88\%) remained per sample. After
removing reads mapping to the host cell genome (\emph{Chlorocebus
sabeus} ), a median of 3.4x10\textsuperscript{6} reads (28\%) remained.
A median of 4.4x10\textsuperscript{5} reads mapped to the DDVax
reference sequence, producing a median coverage of 6055x across all
viral segments (Figure S2). SNVs and short insertion and deletions were
quantified using lofreq, and larger structural variants, including
possible DVGs, were quantified using DI-tector (Beauclair et al., 2018;
Wilm et al., 2012).

The virus remained generally stable over passage in Vero cells. We
report variants with [?] 3\% frequency in coding sequences (Table 1).
Three single nucleotide variants in the glycoprotein precursor gene rose
to above 50\% frequency by P5 (Table 1). A variant at position 31
(predicted to produce the amino acid change G3E in the DDVax NSm-deleted
glycoprotein, equivalent to Gly 133 in the RVFV NSm/Gn/Gc polyprotein,
NC\_014396) rose to 54\% frequency by P5. A variant at position 499
(G159D, equivalent to Gly 289 in ZH501) rose to 55\% by P5. And a
variant at position 926 (N301K, equivalent to Asn 431 in ZH501) rose to
90\% frequency by P5.The highest frequency L segment variant was a
synonymous variant at position 4665 that rose to 16\% by P5. No variants
on the S segment rose above 3\% frequency in any sample. Lofreq did not
identify any short insertion or deletion variants above 3\%. Similarly,
DI-tector did not identify any structural variants (larger insertions,
deletions, incomplete transcripts consistent with DVGs or copy-back
variants) with a frequency [?] 3\%.

\subsubsection*{3.2 Mosquito vector
competence}

{\label{mosquito-vector-competence}}

To measure differences in viral infection kinetics, \emph{Ae.
aegypti}and \emph{Cx. tarsalis} were challenged with 1:1 mixtures of
blood and freshly grown DDVax and then compared against those infected
with MP-12 or the ZH501 parental strain. Because of the need to use
freshly-grown virus for infections, it was not possible to control for
differences in bloodmeal titers. Mean bloodmeal titers ranged from
\textasciitilde{}8.1 logs/ml with DDVax to 6.5 or 6.8
log\textsubscript{10} PFU/ml in MP-12 and ZH501, respectively (Figure
1A). Thus, DDVax titers were significantly higher than that of the other
two virus strains (ANOVA, p=1.8e-5). Virus infection phenotypes were
measured by detection of viral RNA in \emph{Cx. tarsalis} bodies,
legs/wings and saliva at 14 days post-infection (Figure 1B, Table S1).
All saliva samples were also subjected to plaque assay for detection of
infectious virus.

The percentage of \emph{Culex} mosquito DDVax viral RNA positive bodies
was not statistically different from MP-12 or ZH501 infections (Figure
1B, Table S1). However, the RNA genome copy number in \emph{Culex}
or\emph{Aedes} infected with DDVax was at least two
log\textsubscript{10}values lower than those infected with either MP-12
or ZH501 strains, despite exposure of mosquitoes to a DDVax titer over
one log\textsubscript{10} PFU greater than controls (Figure 2).
Dissemination of DDVax viral RNA to \emph{Culex} legs/wings was also
significantly reduced compared to MP-12 (\selectlanguage{greek}χ\selectlanguage{english}\textsuperscript{2}
test,\emph{p} = 2.078e-07). Moreover, infectious DDVax was detected in
only one of 140 \emph{Culex} saliva samples at 14 dpi, whereas 96\% and
82\% of MP-12 and ZH501 infected saliva samples, respectively, showed
CPE consistent with the presence of infectious virus (Table 2,
\selectlanguage{greek}χ\selectlanguage{english}\textsuperscript{2} test, \emph{p} = 2.2e-16 vs MP-12, 2.2e-16, vs
ZH501). To rule out the possibility that sample freeze-thaw compromised
virus viability, an additional subset of saliva samples from 14 dpi
DDVax exposed mosquitoes were assessed for the presence of infectious
virus; still, none was detected (Table 2).\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image1/image1}

\end{center}
\end{figure}

\textbf{Figure 1. Viral RNA detection in RVFV DDVax, MP-12 and
ZH501\emph{in vitro} challenged mosquito bodies, legs/wings and salivary
expectorants at 14 dpi.} Sample positivity rates are listed in S1 Table.
Viral copy number was calculated using a standard curve of diluted L
segment transcripts amplified from a plasmid. Profiles from 3 biological
replicates were combined, with approximately 40 mosquitoes per
replicate.

\emph{Ae. aegypti} from the \emph{in vitro} virus exposure experiments
also showed significantly reduced dissemination in DDVax-infected
mosquitoes compared to those challenged with MP-12 or ZH501,
respectively (\selectlanguage{greek}χ\selectlanguage{english}\textsuperscript{2} test, vs MP-12 \emph{p} = 0.02, vs
ZH501 \emph{p} = 2.2e-16), as indicated by the presence of viral RNA in
legs/wings (Figure 1B). \emph{Aedes aegypti} mosquitoes exposed to DDVax
had no evidence of infectious virus in expectorated saliva, whereas 16\%
and 27\% of saliva samples were CPE-positive in MP-12 and ZH501 infected
mosquitoes, respectively (Table 2, \selectlanguage{greek}χ\selectlanguage{english}\textsuperscript{2} test, vs
MP-12\emph{p} = 2.2e-16, vs ZH501 2.821e-09).

\textbf{3.3 Dose response curve}

We expected that DDVax would not be found at significant levels outside
mosquito midguts, as described in previous reports of plaque assays for
infectious virus(Crabtree et al., 2012). However, our challenge
experiments showed unexpectedly high levels of DDVax RNA-positive,
CPE-negative saliva samples (Table 2, Table S1, Figure 1B). We
hypothesized that the high levels of DDVax viral RNA in saliva may have
been due to the high viral titer of the infectious bloodmeal (Figure
1A), which could have overwhelmed natural infection barriers. Therefore,
to confirm that viral RNA positivity varied as a function of bloodmeal
titer, a second DDVax challenge was performed with \emph{Cx.
tarsalis}mosquitoes, using virus serial dilutions. Bloodmeals containing
6.2, 4.5, and 3.5 log\textsubscript{10} PFU/ml DDVax were provided.
There was a trend for reduction of viral RNA in bodies, legs/wings, and
saliva samples as the bloodmeal titer decreased (Table S2, Figure S3).
However, strikingly, there was still detectable viral RNA in salivary
expectorants with all viral dilutions, including the 3.5
log\textsubscript{10} PFU/ml virus meal.

\textbf{3.4 Mosquito challenge on inoculated goats}

To further test the environmental safety profile of DDVax, goats were
inoculated with either DDVax or MP-12 viruses. Mosquitoes were allowed
to directly feed on the goats at 1 and 2 days post-inoculation (Figure
S1). On day 3, blood was collected into EDTA-tubes and transferred to
water-jacketed feeders for mosquito challenge in the laboratory. Numbers
of engorged mosquitoes from each daily goat feeding are listed in Table
S3. Sera from all goat blood specimens were negative for DDVax or MP-12
by plaque assay at 1, 2, and 3 dpi (limit of detection 1
log\textsubscript{10} PFU/ml). However, trace levels of viral RNA were
detectable by RT-qPCR (Figure S4). After a 7-day extrinsic incubation
period, \emph{Aedes} and \emph{Culex} bodies showed evidence of
infectious MP-12 by plaque assay (Figure 2), indicative of midgut
infections, as previously described(Crabtree et al., 2012; Kading et
al., 2014). Viral prevalence was highest in \emph{Aedes} (28\%) exposed
to goats at 1 day post vaccination with MP-12 strain; these
\emph{Aedes}mosquito infection rates decreased to 12\% and 6\% in
mosquitoes that fed on goats 2 and 3 days post-vaccination,
respectively. In contrast, 6\% (day 1), 2\% (day 2) and 5\% (day 3) of
\emph{Aedes} mosquitoes that fed on DDVax-inoculated goats were positive
for infectious virus by CPE assay after a 7-day incubation period.
Across the time series,\emph{Aedes} mosquitoes exposed to MP-12
vaccinated goats showed significantly higher rates of virus-positive
bodies than those exposed goats inoculated with DDVax
(\selectlanguage{greek}χ\selectlanguage{english}\textsuperscript{2} test, \emph{p} = 0.011). \emph{Culex} showed low
rates of MP-12 virus infection (\emph{\textless{}} 10\%) and no evidence
of infection with DDVax. Four of 87 \emph{Culex} mosquitoes that fed on
goats vaccinated with MP-12, and 0/59 \emph{Culex} mosquitoes that fed
on goats inoculated with DDVax, showed evidence of infection after a
7-day incubation. The differences in \emph{Culex} were not significant,
possibly due to low feeding rates (Table S3). All mosquito bodies that
were CPE-positive were assessed for the presence of disseminated live
virus in legs/wings. However, none of the mosquitoes that became
infected after feeding on inoculated goats showed evidence of infectious
virus in disseminated infection (positive legs/wings).

\textbf{Hosted file}

\verb`image2.emf`  available at \url{https://authorea.com/users/435941/articles/538626-human-vaccine-candidate-ddvax-development-against-rift-valley-fever-dissemination-safety-studies-in-mosquitoes}

\textbf{Figure 2. Infectious DDVax or MP-12 detected in bodies from
mosquitoes fed on inoculated goats} . \emph{Aedes} or
\emph{Culex}mosquitoes were fed on goats (n= 3 per virus strain) as
indicated, held for 7 days, then processed for by plaque assay(Table
S3). Graph shows percentage of samples at each day post-inoculation that
were CPE positive, indicative of infectious virus. No infectious virus
was detected in legs/wings samples from virus-positive
mosquitoes.\emph{Aedes} DDVax, n= 50, 50 and 98 for days 1, 2, 3
respectively.\emph{Aedes} MP-12, n= 64, 60 and 100 for days 1, 2, 3
respectively.\emph{Culex} DDVax, n= 11, 8 and 40 for days 1, 2, and 3,
respectively, and showed no evidence of infectious virus at any
timepoint.\emph{Culex} MP-12, n= 22, 9 and 56 for days 1, 2, 3
respectively.

\textbf{3.5 Viral growth curves in mosquito cell lines}

To further characterize DDVax replication kinetics compared to MP-12 and
ZH501 strains, growth curves were performed in three insect cell lines.
Aag2 (\emph{Ae. aegypti,} embryonic), ATC10 (\emph{Ae. aegypti,} larval)
and Ct (\emph{Cx. tarsalis,} embryonic) cells were infected with DDVax,
MP-12 or ZH501 over 6-day time courses. The \emph{Aedes aegypti} larval
cell line ATC10 was not susceptible to infection with any virus strain.
DDVax replicated in Aag2 cells to lower peak titers than did MP-12 or
ZH501 strains (Figure S5) (random effects mixed model ANOVA, p =
8.0e-4). Similarly, DDVax also attained lower titers than control
viruses in Ct cells (random effects mixed model ANOVA, \emph{p} =
3.5e-4). MP-12 grew to similar peak titers in Ct and Aag2 cells, at 9.1
and 9.5 log\textsubscript{10} PFU/ml, respectively. Peak ZH501 titers
were 8.0 and 6.9 log\textsubscript{10} PFU/ml, in Ct and Aag2
cells\emph{,} respectively. The virulent strain caused syncytial
formation and lifting of cell monolayers, consistent with
pathogenicity(Turell, Gargan, \& Bailey, 1984), which could have
affected final titers. Lastly, mean peak DDVax titers were 7.1 and 6.3
log\textsubscript{10} PFU/ml, in Ct and Aag2 cells\emph{,} respectively,
which are lower than peak titers for MP-12. DDVax grew better in Ct
cells than in Aag2 cells (Two way ANOVA, \emph{p} = 4.5e-5), consistent
with the mosquito data. While the calculated DDVax MOI was 0.01 for all
cell lines, the actual MOI was 0.0052 for Aag2, 0.0127 for Ct, and
0.0153 for ATC-10 cells. The difference observed in replication kinetics
of DDVax in the Ct cells as compared with the Aag2 cells may be due in
part to the actual MOI being half the calculated value for Aag2 cells
(Figure S5).

\section*{4 Discussion}

{\label{discussion}}

This study utilized multiple approaches to demonstrate the relative
safety of the DDVax vaccine candidate from the perspective of relevant
mosquito species transmissibility and regulations regarding potential
environmental impacts following field-use. These experiments were
designed as part of a series of safety studies required prior to human
clinical trials. DDVax showed favorable environmental safety profiles
(e.g., low mosquito dissemination, and impaired transmission from
inoculated livestock) compared to MP-12 vaccine and the wild-type
parental virus, ZH501. Mosquitoes in two epidemiologically-relevant
genera were challenged with viral titers up to 2 to 5
log\textsubscript{10} PFU/ml higher than mosquitoes would be expected to
encounter in the field from vaccinated animals, and there was only one
questionably positive transmission event. In a previous study, sheep
vaccinated with DDVax did not develop any detectable vaccine-associated
viremia following inoculation, suggesting that the overall burden of
DDVax in animals is very low (Bird et al., 2011). Additionally, DDVax
viral RNA copy numbers in bodies and legs/wings were significantly
reduced in both \emph{Aedes} and \emph{Culex} compared to those infected
with either MP-12 or ZH501 (Figure 2). This result is consistent with
previously observed impaired viral dissemination phenotype in mosquitoes
due to the deletion of the NSm coding region (Crabtree et al., 2012;
Kading et al., 2014). Only one of 140 mosquito saliva samples contained
live DDVax virus (Table 2), which was also consistent with previous
experiments(Crabtree et al., 2012). This single positive saliva sample
showed a single plaque, which may not have been infectious and for which
we cannot rule out the possibility that it represents low-level
contamination. Expected virus infection rates in these mosquito species
have previously ranged between 63-84\% for virulent recombinant ZH501
(rZH501) for \emph{Ae. aegypti} (Crabtree et al., 2012; Kading et al.,
2014), 58 -- 95\% for \emph{Cx. tarsalis} (Bergren, Borland, Hartman, \&
Kading, 2021; Turell, Wilson, \& Bennett, 2010) with midgut titers of
over 6 log\textsubscript{10} PFU in actively infected \emph{Cx.
tarsalis} (Bergren et al., 2021). Similarly, up to 100\% infection
occurred MP-12 infected \emph{Cx.} \emph{pipiens} (Turell \& Rossi,
1991). In contrast, we expected 0\% infection with DDVax
infected\emph{Ae. aegypti} (Crabtree et al., 2012). Overall, we observed
similar results between ZH501 and MP-12 strains, with a significant
reduction in infection of mosquitoes with DDVax.

While DDVax RNA was detectable in multiple body compartments of the
mosquito, infectivity was very reduced or nil given the low RNA copy
number detected in mosquitoes 14 days post \emph{in vitro} infection
(Figure 1B). For example, if mosquitoes imbibed a 5 \selectlanguage{greek}µ\selectlanguage{english}l blood meal of 8
log\textsubscript{10} PFU/ml, then 5.7 log\textsubscript{10} PFU would
have been acquired. In our study, after two weeks incubation, 2.9
log\textsubscript{10} mean RNA copies were detected in
\emph{Culex}bodies, 1.8 log\textsubscript{10} RNA copies in legs/wings
and 1.5 log\textsubscript{10} RNA copies saliva (Figure 1B), suggesting
that the virus may have somehow disseminated and persisted at a low
level, but was not actively replicating in the mosquitoes. By
comparison, mosquitoes of each species exposed to MP-12 and ZH501 had
RVFV RNA copy numbers between 7-8 log\textsubscript{10} by 14 days
post-exposure (Figure 1B) after exposure to a blood meal containing
greater than an order of magnitude less virus than that of DDVax (Figure
1A). This pattern was consistent with the results of the dose response
experiment, in which the RNA copy number in different tissue
compartments seems to be relatively stable after 14 days across all
three exposure doses (Figure S3). It is not clear at this time how this
spread would be occurring. Further, it is expected that RNA copy numbers
would exceed infectious titers rendering the truly infectious virus
population even lower(Wichgers Schreur et al., 2021).

Consistent with these findings, Kading et al.(Kading et al., 2014)
reported 80\% infection and 60\% dissemination rates of rZH501
by\emph{Ae. aegypti} mosquitoes, compared with 0\% infection and 0\%
dissemination rates of the rZH501-delNSm (NSm deletion) strain, by
plaque assay. Nevertheless, in rZH501-delNSm infections, viral protein
was detected in most mosquitoes by immunofluorescence assay (IFA),
consistent with viral protein translation with defective packaging or
dissemination. Moreover, IFA foci in the midguts of mosquitoes infected
with rZH501-delNSm were also very small compared with extensive midgut
foci characteristic of rZH501(Kading et al., 2014). Therefore, detection
of viral RNA (this study) and antigen (Kading et al., 2014) outside the
midgut, in the absence of infectious virus, warrants further study.

Viral RNA detected in \emph{Culex} saliva could be the result of
cell-to-cell spread of DDVax through tissues in the absence of efficient
viral assembly, or possibly ``leakage'' of virions from the alimentary
tract in the absence of viral replication. Romoser and colleagues
reported the particular affinity of virulent ZH501 RVFV for the cardia,
intussuscepted foregut, fat body and salivary glands in \emph{Culex
pipiens} mosquitoes(Lerdthusnee, Romoser, Faran, \& Dohm, 1995; Romoser,
Faran, Bailey, \& Lerdthusnee, 1992). The cardia and intussuscepted
foregut are transitional tissues between the esophagus and the anterior
midgut in the mosquito digestive tract(Romoser et al., 1992). Salivary
glands are proximal to this region, embedded in fat body. One possible
explanation is that DDVax retained similar tissue affinity in the
absence of NSs and NSm, and, when combined with presumed less efficient
viral assembly, led to detection of viral RNA but no infectious virus
(Table 2, Figure 1B, S1 Table). In addition, Romoser et al. reported
that, in \emph{Culex} , RVFV ZH501 was able to escape to peripheral
tissues as early as 1 day following an infectious blood meal (Romoser et
al., 1992), making it particularly rapid in disseminating compared to
other arboviruses, eg., flaviviruses, which often require at least a
week to reach the salivary glands(Sanchez-Vargas et al., 2009),
depending on extrinsic incubation temperature. RVFV affinity for
salivary glands was substantiated by the DDVax dose response experiment,
in which nearly 19\% of mosquitoes showed viral RNA in salivary
expectorants at the lowest bloodmeal titer of 3.5
log\textsubscript{10}PFU/ml (Table S2). This level approached that of
the presence of viral RNA in legs/wings.

To address concern about the one possible transmission event, \emph{Cx.
tarsalis} mosquitoes were subsequently challenged with artificial blood
meals containing a range of viral titers. As expected, the percentage of
mosquitoes that became infected, as determined by RNA genome copy
number, decreased proportionally with the titer of DDVax in the
artificial blood meal, but did not reach zero. The stable persistence of
DDVax RNA in different tissue compartments was evident in all dosing
groups (Figure S3). As experimentally predicted, the higher the blood
meal titer, the higher the percentage of mosquitoes had detectable RNA,
although infectious virus was not assayed in mosquitoes challenged with
lower titer blood meals.

These results were further confirmed and placed into a realistic
epidemiological context by feeding mosquitoes on inoculated goats.
Infection of goats with wild-type ZH501 was not possible in this study
due to biosafety considerations. Mosquitoes were fed on goats on days
1-3 post-inoculation with DDVax or MP-12. As expected, goats did not
develop any detectable viremia, as determined by plaque assay. However,
small ruminants, e.g. sheep, would be expected to develop a viremia
ranging from \textasciitilde{}5-6
log\textsubscript{10}TCID\textsubscript{50}/ml titers between 1-3 days
post infection with a wild-type strain(Wichgers Schreur et al., 2021).
Similarly, neither Wilson et al.(Wilson et al., 2014) nor Nyundo et
al.(Nyundo et al., 2019) observed any detectable viremia in ruminants
following vaccination with MP-12 strain. Morrill et al.(Morrill et al.,
1991) noted a transient, low-titer viremia in lambs vaccinated with
MP-12 strain. Sheep inoculated with DDVax failed to develop any
detectable viremia (Bird et al., 2011). Therefore, it was surprising to
observe that mosquitoes fed on these inoculated goats and held for seven
days post-feeding developed infections (Figure 2, S3 Table).

Analysis on goat serum samples showed very low (\textless{}10 RNA copies
/ml) RNA levels of RVFV in goat serum (Figure S4), which we interpreted
to represent residual, circulating virus as opposed to actively
replicating virus. The sensitivity of mosquito feeding was able to pick
up this residual viral inoculum, however none of these mosquitoes
developed a disseminated infection by 7 days post-exposure. For
infection with ZH501 strain, dissemination has previously been
documented to occur as early as 3 days post exposure(Romoser et al.,
1992), with all mosquitoes having developed a disseminated infection by
10 days post-exposure(Kading et al., 2014).

Mosquito infectivity also becomes a function of volumetric constraints
of mosquito blood meal size. While the probability of one mosquito
imbibing infectious virions is lower at low virus titers, many
mosquitoes imbibing a blood meal simultaneously would draw a larger
collective volume of blood that could result in one or more mosquitoes
picking up infectious virions. For example, detection of virus in a
single mosquito blood meal is limited to titers \textgreater{}3
log\textsubscript{10} PFU/ml serum, (approximately one PFU in one
microliter of serum in a blood meal) (Kading et al., 2014). For a 25\%
probability of detecting virus in a single 2 \selectlanguage{greek}µ\selectlanguage{english}l mosquito blood meal, the
serum titer needs to be 2.72 log\textsubscript{10} PFU/ml (95\% CI
2.19--3.27), while for a 50\% probability of detection, the titer needs
to be 3.64 log\textsubscript{10} PFU/ml (95\% CI 3.20--4.08)(Kading et
al., 2014). Corresponding titers for 75\% and 90\% probabilities of
detection were 4.56 log\textsubscript{10} PFU/ml (95\% CI 4.02--5.10)
and 5.48 log\textsubscript{10} PFU/ml (95\%CI 4.71--6.24),
respectively(Kading et al., 2014).

Wichgers Schreur et al.(Wichgers Schreur et al., 2021) documented the
extraordinary efficiency of RVFV transmission between lambs and\emph{Ae.
aegypti} mosquitoes when using an animal model as opposed to an
artificial system. Approximately 30\% more RVFV saliva-positive
mosquitoes resulted from feeding on viremic lambs than from feeding on a
membrane system(Wichgers Schreur et al., 2021) testifying to the value
of conducting these experiments with an \emph{in vivo} model system to
more realistically represent vertebrate infectiousness to mosquitoes.
While dissemination of DDVax after our 7-day time point cannot be ruled
out, our collective results suggest that transmission risk would be very
low because any disseminated virions would not be infectious. In
addition, based on previous reports, we expected a low combined
probability of a single mosquito imbibing an infectious virion precisely
after inoculated and an extremely low imbibed virus titer. Moreover,
impaired dissemination was due to the deletion of the NSm gene. Finally,
we saw the lack of infectious DDVax expectorated in mosquito saliva even
after a high titer virus challenge. These combined features combine to
give DDVax a safe environmental profile.

4.2 \textbf{Conclusion}

Due to the double gene deletion of NSs and NSm, DDVax has less efficient
viral replication in mosquitoes than a previous vaccine strain, MP-12 or
wild-type ZH501. Mosquitoes were able to imbibe and harbor infectious
DDVax following a high titer challenge in the lab or feeding on
inoculated goats. However, DDVax replication and dissemination was
impaired in mosquitoes, and only one individual mosquito had one DDVax
plaque in its saliva after a high titer challenge. Given the combined
probability of a single mosquito imbibing an infectious virion precisely
after inoculation, the extremely low imbibed virus titer, the impaired
dissemination in mosquitoes due to the deletion of the NSm gene, and the
lack of infectious DDVax expectorated in mosquito saliva even after a
high titer virus challenge, the transmission and dissemination of DDVax
by mosquitoes from vaccinated individuals in an epidemiologically
relevant scenario is highly unlikely.

\section*{Acknowledgements}

{\label{acknowledgements}}

We thank the following for their contributions to this work. Aaron
Brault and Sean Masters at the Centers for Disease control and
Prevention shared the Ct cell line and \emph{Cx. tarsalis} colony,
respectively. Stephanie Porter, Izabela Regan, and Michelle Savran
assisted with goat studies. Marylee Kapuscinski and Mikaela Samsel
assisted with sequencing.

\section*{Ethics Statement}

{\label{ethics-statement}}

The authors confirm that the ethical policies of the journal, as noted
on the journal's author guidelines page, have been adhered to and the
appropriate ethical review committee approval has been received. The US
National Institutes of Health guidelines for the Care and Use of
Laboratory Animals were followed.

\section*{Conflict of Interest}

{\label{conflict-of-interest}}

The authors declare they have no conflict of interest.



\section*{Supporting Information}

{\label{supporting-information}}

Figure S1. Vaccinated goats were exposed to mosquitoes.

Figure S2. Depth of coverage across DDVax segments. Coverage depth was
tabulated using samtools and depth in 10 nt windows is plotted.

Figure S3. Detection of DDVax viral RNA following exposure to limiting
dilutions of virus at 14 dpi.

Figure S4. Viral RNA detected in goat blood at 1, 2, 3 days
post-vaccination.

Figure S5. Mosquito cell culture shows reduced efficiency of viral
replication for DDVax.

Table S1. RNA positive RVFV-exposed samples from \emph{in vitro}mosquito
challenge.

Table S2. RNA positive RVFV DDVax-exposed samples from virus serial
dilution experiment.

Table S3. Proportion of mosquitoes with viral CPE in bodies at 7 days
post-goat-exposure.

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\textbf{Table 1: Single Nucleotide Variants.}\selectlanguage{english}
\begin{longtable}[]{@{}llllllllll@{}}
\toprule
\textbf{Segment} & \textbf{Position (nt)} & \textbf{Coding impact} &
\textbf{Reference Base} & \textbf{Variant Base} & \textbf{P1} &
\textbf{P2} & \textbf{P3} & \textbf{P4} & \textbf{P5}\tabularnewline
\midrule
\endhead
L & 4665 & E1549E & A & G & 0.00 & 0.07 & 0.09 & 0.12 &
0.16\tabularnewline
L & 5483 & K1822M & A & T & 0.04 & 0.03 & 0.03 & 0.03 &
0.04\tabularnewline
L & 5488 & D1824Y & G & T & 0.07 & 0.07 & 0.07 & 0.07 &
0.07\tabularnewline
L & 5513 & R1832I & G & T & 0.00 & 0.00 & 0.00 & 0.00 &
0.03\tabularnewline
L & 6113 & Y2032C & A & G & 0.05 & 0.05 & 0.05 & 0.04 &
0.03\tabularnewline
M & 31\textsuperscript{a} & G3E & G & A & 0.00 & 0.00 & 0.00 & 0.25 &
0.54\tabularnewline
M & 32 \textsuperscript{a} & G3G & G & A & 0.00 & 0.00 & 0.07 & 0.20 &
0.19\tabularnewline
M & 190 & G56E & G & A & 0.00 & 0.00 & 0.00 & 0.00 & 0.09\tabularnewline
M & 300 & K93E & A & G & 0.00 & 0.00 & 0.00 & 0.03 & 0.05\tabularnewline
M & 457 & Y145C & A & G & 0.00 & 0.00 & 0.00 & 0.06 &
0.00\tabularnewline
M & 462 & L147I & C & A & 0.00 & 0.06 & 0.09 & 0.09 &
0.00\tabularnewline
M & 499 & G159D & G & A & 0.00 & 0.00 & 0.00 & 0.11 &
0.55\tabularnewline
M & 808 & R262K & G & A & 0.00 & 0.07 & 0.07 & 0.09 &
0.00\tabularnewline
M & 818 & K265N & A & T & 0.00 & 0.00 & 0.00 & 0.00 &
0.04\tabularnewline
M & 925 \textsuperscript{a} & N301I & A & T & 0.00 & 0.00 & 0.03 & 0.03
& 0.00\tabularnewline
M & 926 \textsuperscript{a} & N301K & T & A & 0.06 & 0.34 & 0.40 & 0.49
& 0.90\tabularnewline
M & 1240 & D406G & A & G & 0.04 & 0.00 & 0.04 & 0.06 &
0.00\tabularnewline
M & 1473 & P484S & C & T & 0.00 & 0.00 & 0.00 & 0.07 &
0.00\tabularnewline
M & 2480 & R819R & G & A & 0.00 & 0.00 & 0.00 & 0.00 &
0.05\tabularnewline
\bottomrule
\end{longtable}

The variants at positions 31 and 32, and those at positions 925 and 926
are not linked.

\textbf{Table 2 Proportion of mosquitoes with infectious virus in saliva
following exposure to RVFV with an artificial membrane feeder.}\selectlanguage{english}
\begin{longtable}[]{@{}llll@{}}
\toprule
\textbf{Species} & \textbf{Virus} & \textbf{\# samples} & \textbf{Saliva
CPE positive (\%)}\tabularnewline
\emph{Ae. aegypti} & DDVax & 120 & 0 (0\%)\tabularnewline
& MP-12 & 120 & 19 (16\%)\tabularnewline
& ZH501 & 118 & 32 (27\%)\tabularnewline
\emph{Cx. tarsalis} & DDVax & 120 & 1 (\textless{} 1\%)\tabularnewline
& DDVax & 20* & 0(0\%)\tabularnewline
& MP-12 & 120 & 115 (96\%)\tabularnewline
& ZH501 & 110 & 90 (82\%)\tabularnewline
& ZH501 & 15* & 10 (67\%)\tabularnewline
\bottomrule
\end{longtable}

*, no freeze-thaw\selectlanguage{english}
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\includegraphics[width=0.70\columnwidth]{figures/Fig2/Fig2}

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