Vaccine strains of Rift Valley fever virus exhibit attenuation at the maternal–fetal placental interface

Cynthia M McMillen; Christina Megli; Rebecca Radisic; Lauren B Skvarca; Ryan M Hoehl; Devin A Boyles; Jackson J McGaughey; Brian H Bird; Anita K McElroy; Amy L Hartman

doi:10.1128/jvi.00983-24

. 2024 Jul 17;98(8):e00983-24. doi: 10.1128/jvi.00983-24

Vaccine strains of Rift Valley fever virus exhibit attenuation at the maternal–fetal placental interface

Cynthia M McMillen ^1,^2,^✉, Christina Megli ³, Rebecca Radisic ⁴, Lauren B Skvarca ⁵, Ryan M Hoehl ¹, Devin A Boyles ¹, Jackson J McGaughey ¹, Brian H Bird ⁴, Anita K McElroy ^1,⁶, Amy L Hartman ^1,^2,^✉

Editor: Stacey Schultz-Cherry⁷

¹University of Pittsburgh, Center for Vaccine Research, Pittsburgh, Pennsylvania, USA

²Department of Infectious Diseases and Microbiology, University of Pittsburgh, School of Public Health, Pittsburgh, Pennsylvania, USA

³Division of Maternal-Fetal Medicine, Division of Reproductive Infectious Disease, Department of Obstetrics, Gynecology and Reproductive Sciences, University of Pittsburgh School of Medicine and the Magee-Womens Research Institute, Pittsburgh, Pennsylvania, USA

⁴One Health Institute, School of Veterinary Medicine, University of California, Davis, California, USA

⁵Department of Pathology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania, USA

⁶Department of Pediatrics, Division of Pediatric Infectious Disease, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania, USA

⁷St. Jude Children's Research Hospital, Memphis, Tennessee, USA

^✉

Address correspondence to Amy L. Hartman, hartman2@pitt.edu

^✉

Address correspondence to Cynthia M. McMillen, cmm287@pitt.edu

B.H.B. is an inventor of patents describing the development of the ΔNSs-ΔNSm-rZH501 vaccine candidate technology (USPTO: 8,673,629, 9,439,935, and 10,064,933). The remaining authors declare that they have no competing interests.

Roles

Cynthia M McMillen: Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Visualization, Writing – original draft, Writing – review and editing

Christina Megli: Conceptualization, Data curation, Investigation, Methodology, Writing – review and editing

Rebecca Radisic: Formal analysis, Investigation, Methodology, Visualization

Lauren B Skvarca: Formal analysis, Visualization, Writing – review and editing

Ryan M Hoehl: Data curation, Investigation, Methodology

Devin A Boyles: Data curation, Investigation, Methodology, Visualization

Jackson J McGaughey: Data curation, Investigation, Methodology

Brian H Bird: Formal analysis, Methodology, Supervision, Visualization, Writing – review and editing

Anita K McElroy: Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Resources, Validation, Visualization, Writing – review and editing

Amy L Hartman: Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Supervision, Validation, Visualization, Writing – original draft, Writing – review and editing

Stacey Schultz-Cherry: Editor

PMCID: PMC11334480 PMID: 39016561

ABSTRACT

Rift Valley fever virus (RVFV) infection causes abortions in ruminant livestock and is associated with an increased likelihood of miscarriages in women. Using sheep and human placenta explant cultures, we sought to identify tissues at the maternal–fetal interface targeted by RVFV. Sheep villi and fetal membranes were highly permissive to RVFV infection resulting in markedly higher virus titers than human cultures. Sheep cultures were most permissive to wild-type RVFV and ΔNSm infection, while live-attenuated RVFV vaccines (LAVs; MP-12, ΔNSs, and ΔNSs/ΔNSm) exhibited reduced replication. The human fetal membrane restricted wild-type and LAV replication, and when infection occurred, it was prominent on the maternal-facing side. Type I and type III interferons were induced in human villi exposed to LAVs lacking the NSs protein. This study supports the use of sheep and human placenta explants to understand vertical transmission of RVFV in mammals and whether LAVs are attenuated at the maternal–fetal interface.

IMPORTANCE

A direct comparison of replication of Rift Valley fever virus (RVFV) in sheep and human placental explants reveals comparative efficiencies and permissivity to infection and replication. Vaccine strains of RVFV demonstrated reduced infection and replication capacity in the mammalian placenta. This study represents the first direct cross-host comparison of the vertical transmission capacity of this high-priority emerging mosquito-transmitted virus.

KEYWORDS: Rift Valley fever, bunyavirus, phlebovirus, pregnancy, placenta, sheep, human, vertical transmission

INTRODUCTION

Rift Valley fever (RVF) is a zoonotic, mosquito-transmitted disease in Africa and the Middle East that primarily affects domesticated livestock (sheep, goats, cattle, and camels) and can cause mild to severe disease in humans. During rainy seasons, when mosquito populations expand, outbreaks are accompanied by high rates of vertical transmission in livestock, leading to fetal death. An entire generation of sheep can be lost in a single outbreak in part due to these “abortion storms,” causing devastating economic impacts on affected regions (1). Abortions in livestock are associated with diffuse placental hemorrhage and necrosis (2) along with cerebral and musculoskeletal deformities in fetuses (3, 4). Given its known effects on pregnant animals, Rift Valley fever virus (RVFV) infection might also cause miscarriages in women (5). A study in Sudan showed that women infected with RVFV during pregnancy are approximately four times more likely to have a “late miscarriage,” defined as a second- or third-trimester fetal loss. Cases of late gestation fetal infection, including infants born at full term with RVF disease, have been documented in humans (6, 7).

Live-attenuated vaccines (LAVs) are currently used in endemic areas to minimize the incidence of disease in livestock (8, 9); however, the residual virulence of some of these attenuated strains may still lead to adverse events in pregnant animals, including fetal loss or deformity (10, 11). Attenuated vaccine strains of RVFV were generated through serial passaging in vivo, chemical mutagenesis, or by mutating or removing one or both virulence factors, NSs or NSm. Two LAV strains, Smithburn and Clone 13 (similar to ΔNSs), are regularly used in livestock throughout Africa (8, 9). Both strains retain some virulence, which can lead to fetal death or teratogenic events in pregnant livestock; therefore, pregnant animals are not routinely vaccinated, leaving them vulnerable to infection by virulent field strains of RVFV (10, 11). Other LAVs were developed to overcome the limitations of Smithburn and Clone 13, including MP-12, an attenuated strain generated by chemical mutagenesis that resulted in nine amino acid mutations across all three segments of the RNA genome (12, 13). MP-12 has undergone early-stage clinical trials in humans (14, 15). There are conflicting reports on the safety of MP-12 during pregnancy. Sheep vaccinated with high doses of MP-12 during early gestation delivered deformed fetuses (16), whereas those vaccinated with a lower dose during late gestation delivered healthy lambs (17, 18). A next-generation LAV strain made using reverse genetics to delete both NSs and NSm (ΔNSs/ΔNSm) appeared to be safe in pregnant sheep, even when vaccinated during early gestation (19). Although there are no approved RVFV vaccines for humans, clinical trials are underway with updated versions of ΔNSs/ΔNSm and several other promising candidates (20 –22). Identifying a safe and effective vaccine to protect humans from severe RVF will be a tremendous benefit to the field. However, early-stage human clinical trials are unlikely to provide sufficient safety data in pregnant women as they are generally excluded from these trials. Moreover, as RVFV is particularly pathogenic to pregnant mammals, it is critical to understand the pathogenic mechanisms at the maternal–fetal interface.

Despite the well-documented outcomes of RVFV fetal infection in ruminant livestock, it is not clear whether the seemingly lower rates of miscarriages reported in humans are due to limited epidemiologic data leading to reporting bias or biological resistance mechanisms impeding human fetal infection. Although sheep and humans are eutherian mammals utilizing placental membranes for fetal development, each has significant differences in cellular composition, blood flow patterns, and tissue structures composing the placenta (23, 24). Ex vivo systems using placenta explants have provided important information regarding the pathologies and immune responses associated with other congenital infections (25, 26). Here, we use placenta explants to compare the permissivity of the maternal–fetal interface of sheep and humans to infection with RVFV. Furthermore, we use the explants to screen the replicative capacity of LAVs, given their variable abilities to cause abortions and teratogenic effects in livestock in vivo (10, 11, 16 –19). Since limited epidemiological data exist to fully understand the impact of vertical transmission of RVFV in pregnant women, ex vivo analyses can play a pivotal role in risk assessment.

RESULTS

To provide a relevant comparison between human and sheep placenta with regard to the permissivity of semi-intact structures located at the maternal–fetal interface, it is important to delineate the structural differences between the placenta of humans and sheep. Unlike human placentas, which are discoidal, sheep placentas have a cotyledonary structure containing up to 70–100 individual fetal-derived cotyledons that interdigitate with maternal caruncle microvilli constituting what are known as placentomes (Fig. 1A). Villi, which are tree-like branches made of trophoblasts and fetal vasculature, play an important role in nutrient and oxygen exchange between the fetus and the mother. The placentomes are distributed across the entire gestational sac and connected by the allantoic and chorionic membranes (27). The fetal membranes provide a protective barrier between the fetus and the mother. Each placentome fits inside a specialized pocket within the maternal decidua, called a caruncle. The trophoblast-lined cotyledon connects with the decidua through weak invasion by cytotrophoblasts. The cytotrophoblasts fuse with, or invade into, the uterine epithelial cells, creating an epitheliochorial barrier that defines livestock placenta.

Humans, in contrast, have a single, discoid placenta (Fig. 1B). Like sheep, villi of the human placenta contain fetal vasculature and are lined by trophoblast. Unlike sheep, trophoblasts lining human placental villi are bathed in flowing maternal blood, which is critical to providing nutrients and oxygen to the fetus. The chorioamniotic membrane surrounds the developing fetus, maintains the amniotic fluid, protects the fetus from injury, and regulates fetal temperature. Decidualized endometrium at the implantation site, known as the decidua basalis, comes in direct contact with villi via extravillous trophoblasts that invade the endomyometrium to establish a vascular flow to the developing placenta. Blood flows from adapted uterine vessels in the decidua into the intervillous space.

Separation and dissection of livestock placenta into villi, chorionic membrane, and allantoic membrane (28 –30) and human placenta into villi, fetal membranes, and decidua are feasible (25, 31) (Fig. 1C). Maternal decidual tissues are generally not available from sheep after natural birth because they fully separate from the placenta at delivery and, thus, are not analyzed in this study. We hypothesize that culturing placental tissue sub-types will allow the identification of regions of the maternal–fetal interface that are susceptible to RVFV infection.

Sheep placenta explants are permissive to infection with RVFV vaccine strains

Term placentas and one preterm placenta were collected from four sheep housed at a local farm in western Pennsylvania (Table 1). Dissected villi, chorionic membranes, and allantoic membranes were inoculated with 1 × 10⁵ plaque-forming units (pfu) of wild-type RVFV (ZH501) or attenuated strains (ΔNSm, ΔNSs, MP-12, or ΔNSs/ΔNSm) (Fig. 2A and B). Culture supernatant was collected every 12–24 hours to generate a viral replication curve by quantitative real-time PCR (q-RT-PCR) (Fig. 2C).

TABLE 1.

Sheep placenta demographics^a

Sheep	Breed	Gestation	Offspring sex
Donor 1	Texel x Suffolk	Approx. 4 mo	–
Donor 2^b	Texel x Suffolk	Term	Two males
Donor 3	Texel x Dorset	Term	Male
Donor 4^b	Texel x Suffolk	Term	Two males

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^{^a}

x, crossed breed; –, unknown.

^{^b}

Twins.

Villi and both fetal membranes obtained from sheep were permissive to infection with all strains of RVFV (Fig. 2C). Viral RNA was detected as early as 24 hpi and increased through 72 hpi. RVFV ZH501 and ΔNSm produced the highest levels of viral RNA in the culture supernatant compared to the other strains, with some cultures reaching as high as 5 × 10⁸ pfu equivalents/mL (Fig. 2C). For wild-type ZH501, there was a steady 3-log increase in virus over 72 hours for all tissues, whereas the replication curve for the villous and chorionic membrane cultures infected with ΔNSm showed a biphasic response. After infection, villi consistently shed more RVFV ZH501 and ΔNSm compared to the allantoic and chorionic membranes. The strains with the lowest tissue replication in sheep explants were ΔNSs, MP-12, and ΔNSs/ΔNSm (approx. 1 × 10⁴ - 1 × 10⁶ pfu equivalents/mL), and replication appeared to plateau after 24 hours post-infection (Fig. 2C; Fig. S1).

Consistent with the q-RT-PCR results, immunohistochemistry revealed widespread infection within the cytoplasm of cells throughout the villi and allantoic and chorionic membranes inoculated with ZH501 as depicted by moderate levels of RVFV nucleoprotein immunopositivity (Fig. 3; Table S1). Immune cells with detectable viral nucleoprotein were also observed in the stroma of the tissues infected with ZH501. For ΔNSm, strong nucleoprotein immunopositivity was present in the allantoic membrane and villous in addition to stromal immune cells. The presence of nucleoprotein in the chorionic membrane was inconclusive due to the artifact of background reactivity. For the other attenuated RVFV strains, moderate levels of cytoplasmic and membranous viral antigen were detected in the chorionic membrane inoculated with MP-12, and immune cells containing nucleoprotein were rarely identified in the stroma and trophoblasts. Low levels of immunopositivity were observed in the chorionic membrane, and none was detected in the allantoic membrane of tissues exposed to ΔNSs/ΔNSm in culture. In contrast, strong nucleoprotein deposition was seen in the cytoplasm of cells within the villous, and RVFV antigen-positive immune cells were also observed in the stroma of the villous of ΔNSs/ΔNSm-infected tissues. ΔNSs, however, did not appear to infect sheep placenta explant cultures as negligible antigen staining was identified. Overall, ZH501 and ΔNSm caused widespread infection of the fetal membranes and villous. MP-12 and ΔNSs/ΔNSm infect the chorionic membrane and/or villous to varying degrees.

Fig 3 — ZH501 and ΔNSm undergo widespread infection in sheep placenta explants. Immunohistochemistry (IHC) images of sheep allantoic membrane (AM), chorionic membrane (CM), and villi (V) cultures infected with RVFV ZH501, ΔNSm, ΔNSs, MP-12, and ΔNSs/ΔNSm or uninfected controls. (A) Images were taken at 20× magnification by light (optical) microscopy. (B) 40× magnification images of sheep placenta cultures infected with MP-12. Red-brown = RVFV nucleoprotein. Blue staining = cell structural counterstain. Black arrowheads highlight regions infected with RVFV.

All vaccine strains infect human villous explants, but tissue distribution differs in viruses lacking the viral NSs protein

Human placentas from livebirths were collected from women near term (32- and 36-week gestation) (Table 2). Dissected decidua, fetal membranes, and villi were inoculated with 1 × 10⁵ pfu of RVFV ZH501, ΔNSm, ΔNSs, MP-12, or ΔNSs/ΔNsm, and culture supernatant was collected over time to generate a viral replication curve (Fig. 4A and B).

TABLE 2.

Human placenta demographics

Human	Gestation	Offspring sex	Notes
Donor 1	35 weeks, 6 days	Female	Peripheral biopsy of placenta accreta specimen
Donor 2^a	32 weeks, 4 days	Female	Dichorionic, diamniotic twins Suspected intrauterine growth restriction (male)
Donor 3^a	32 weeks, 4 days	Male

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^{^a}

Twins.

Fig 4 — Human placenta explant cultures are permissive to RVFV strains. (A) Representative image of a whole (top) and dissected (bottom) human placenta and decidua. (B) Experimental timeline for human placenta explant culture infections. Tissue dissections were inoculated with 1 × 10⁵ pfu RVFV ZH501, ΔNSs, ΔNSm, MP-12, and ΔNSs/ΔNSm for 1 hour (n = 3–5 each). Virus was removed and washed prior to the addition of culture growth medium. Culture supernatant was collected at 0, 24, 36, 48, and 72 hpi. (C) q-RT-PCR was performed to quantitate virus production over time. Dashed line = LOD. Statistical significance was determined by one-way ANOVA compared to ZH501 cultures. ^*P < 0.05, ^**P < 0.01, ^***P < 0.001, and ^****P < 0.0001. n.s. = not significant. Red, green, purple, and orange asterisks designate the P-value of ΔNSs, ΔNSm, MP-12, and ΔNSs/ΔNSm compared to ZH501, respectively.

Like the sheep cultures, ZH501 and ΔNSm generated the highest levels of viral RNA over time, with some human explant cultures reaching 10⁶ pfu equivalents/mL. ΔNSs and MP-12 reached just above 10⁵ pfu equivalents/mL in villous cultures by 72 hours (Fig. 4C; Fig. S2). Surprisingly, villous and decidua cultures with the double mutant virus, ΔNSs/ΔNSm, reached similar levels of viral RNA as the ZH501 cultures by 72 hours. Overall, virus within villi and decidua replicated at greater efficiency than the fetal membrane as demonstrated by high viral RNA. Fetal membrane cultures reached approximately 5 × 10⁴ pfu equivalents/mL at the endpoint, whereas villi and decidua produced up to 7 × 10⁵ pfu equivalents/mL of viral RNA.

Virus was detected in human villous tissue inoculated with ZH501 or attenuated virus strains by immunohistochemical analysis (Fig. 5A). Immunopositivity was observed in a linear pattern in syncytiotrophoblast and/or within villous stromal cells. Although the number of immunopositive villi appeared fewer in MP-12 and ΔNSs/ΔNSm samples compared to ZH501, additional data are needed to quantify differences in infection kinetics. Immunopositivity was not observed in uninfected villous controls (Table S2).

In contrast to villi, human decidua and fetal membrane cultures inoculated with ZH501 or the attenuated strains demonstrated weak and/or variable immunopositivity. One fetal membrane culture infected with wild-type ZH501 displayed moderate to strong staining in the maternal-facing layers (chorionic trophoblast and decidua capsularis; Fig. 5B; Fig. S3A). This result may indicate regional differences in cellular susceptibility to infection across the fetal membrane.

RVFV NSs antagonizes production of both type I and III interferons in human villous cultures

Innate anti-viral responses, such as the expression of type I and type III interferons (IFNs), can impact viral replication and/or dictate pathogenesis during pregnancy (32). Type I interferons are expressed by most cells to provide systemic control of infection, whereas type III interferons are expressed at barrier surfaces, such as placental trophoblasts, to control infections locally (33, 34). Type I interferons primarily provide protection during the first trimester of pregnancy in humans, whereas type III interferons are more highly expressed and contribute to an anti-viral state during the third trimester (35). RVFV NSs is a type I IFN antagonist, and its role at the maternal–fetal interface has not been evaluated. Furthermore, the antagonistic properties of NSs have not been evaluated in the context of type III IFN. To determine whether there were strain-specific differences in the expression of IFN-α or IFN-λ1, enzyme-linked immunosorbent assays (ELISAs) were performed to quantify protein levels within endpoint culture supernatant. Wild-type ZH501 did not induce IFN-α or IFN-λ1 expression in human placental samples (Fig. 6). In contrast, for attenuated strains lacking NSs expression (ΔNSs and ΔNSs/ΔNSm), IFN-α was elevated in the supernatant of human villous and decidua cultures, with higher concentrations in the villi compared to the decidua. IFN-λ1 is elevated in the supernatant of villous cultures infected with ΔNSs, MP-12, and ΔNSs/ΔNSm compared to uninfected controls, which had undetectable levels of IFN-λ1 (Fig. 6B). IFN-λ1 expression was relatively low or not detected in decidua and fetal membrane cultures infected with LAVs. Overall, viruses without the NSs protein induce higher levels of type I and III interferons in human villous explant cultures, which suggests that NSs could be an antagonist of both types of IFNs in the placenta.

Fig 6 — Attenuated RVFV LAVs induce type I and III IFNs in human placenta. Protein levels of IFN-α (A) or IFN-λ1 (B) were measured in culture supernatant collected at endpoint (48 or 72 hpi) by ELISA. Culture supernatant was collected from villous, decidua, or fetal membrane cultures inoculated with RVFV ZH501, ΔNSm, ΔNSs, MP-12, and ΔNSs/ΔNSm (n = 3–5). IFN-α LOD = 3.2 pg/mL. IFN-λ1 LOD = 13.72 pg/mL. Statistical significance was determined by two-way ANOVA compared to uninfected cultures. ^*P < 0.05; ^**P < 0.01. n.s. = not significant.

Sheep placenta explant cultures produce more virus than human placenta explant cultures

Overall, sheep placenta cultures produced more virus (2 × 10⁵–5 × 10⁸ pfu equivalents/mL) than human cultures (1 × 10⁴–7 × 10⁵ pfu equivalents/mL) (Fig. 2C and 4C). To evaluate the replication kinetics across viruses and tissues, we calculated the slope of the increase in viral RNA in each culture type by linear regression (Fig. 7). We started at 24 hpi for sheep samples and 36 hpi for human samples due to the delay in replication kinetics (Fig. 2C and 4C). The decidua was only collected from human placentas, and thus, there is not a sheep equivalent for comparison. However, in the human decidua, wild-type ZH501 clearly had the steepest slope (i.e., highest virus replication rate), whereas the LAVs did not produce virus after 36 hpi, as portrayed by the near-zero or slightly negative slope (Fig. 7, left panel). For the fetal membranes (Fig. 7, middle panel), sheep cultures had steeper slopes compared to human cultures across all virus strains. All LAVs had a positive slope in sheep allantoic membrane cultures, whereas ΔNSs and MP-12 did not grow well after 36 hpi (slope: −0.007 and −0.111, respectively) in chorionic membranes. ΔNSs/ΔNSm grew quite well in sheep chorionic membranes. For all strains, including ZH501, the virus replication rates were near zero or negative within human fetal membrane cultures. Villi were highly permissive to virus replication for all strains in both sheep and human samples (Fig. 7, right panel), except for ΔNSs, which did not grow in human villous cultures. The slope of the increase in viral RNA was significantly different (P = 0.197) between human villous cultures infected with ZH501 versus ΔNSs/ΔNSm; all other comparisons between tissues were not significant. Taken together, using the slope of the increase in viral RNA over time provides insight into the relative permissivity of sheep and human maternal–fetal interface structures.

Fig 7 — The rate of virus production is similar between hosts, despite delayed production in human cultures. A simple linear regression was performed to calculate the rate of virus production (slope) from virus growth curves for decidua, fetal membrane, and villous cultures starting at 24 hpi (sheep) and 36 hpi (humans). Positive slope = red shades; no slope = black. Individual slopes are depicted by white numbers.

DISCUSSION

There are multiple mechanisms by which pathogens can undergo vertical transmission, an event in which a fetus is infected through exposure from its infected or colonized mother. TORCH pathogens (Toxoplasma gondii, other, rubella virus, cytomegalovirus, herpes simplex virus) can adversely affect the developing human fetus through several potential mechanisms including transplacental transmission, direct insult to the placenta, fetal–maternal hemorrhage, transmission across fetal membranes, or ascending sexual transmission. Pathogens also induce inflammatory responses at the maternal–fetal interface that adversely affect the fetus (33, 36). The major structures at the maternal–fetal interface that protect against vertical transmission are the maternal decidua and fetal-derived membranes (amnion and chorioallantois) and villi (Fig. 1). For many pathogens, the mechanism of vertical transmission across the maternal and fetal structures is incompletely understood. Although RVFV has caused massive abortion storms in livestock since the year 1930 (37, 38), few studies have examined the cellular tropism at the maternal–fetal interface or the routes that RVFV could take to reach the fetus in utero.

In a previous study, Odendaal et al. examined placentas from naturally aborted sheep during a 2010–2011 RVF outbreak in South Africa (39). Extensive RVFV antigen immunostaining was found within trophoblasts in the cotyledonary chorioallantois. In the intercotyledonary areas, such as the chorioallantoic membrane, RVFV antigen was sparse yet detectable. Significant necrosis was observed in the trophoblasts and endothelial cells of the cotyledonary and intercotyledonary chorioallantois, whereas necrosis of the villous trophoblasts was typically diffuse. In a separate experimental study in pregnant sheep, inoculation with the wild-type RVFV strain 35/74 at one-third gestation resulted in abortion at 6 dpi, and immunostaining showed widespread infection of maternal epithelial cells and fetal trophoblasts lining the hematophagous zone of the placenta (2). Placentas showed signs of hemorrhage of the maternal villi, extensive necrosis of the maternal epithelium, and necrosis of the fetal trophoblasts; analyses of the fetal membranes were not performed in this study. These two studies provide important insights into RVFV infection of the maternal–fetal interface of livestock. However, experimental studies in pregnant sheep are expensive and cumbersome and require levels of agricultural animal biocontainment that are restrictive. Thus, given the limited data from both natural and experimental RVFV infection during pregnancy in sheep, we sought to develop tractable placental explant cultures for direct comparison with their human counterparts.

There is increasing clinical evidence of vertical transmission of RVFV during human pregnancy and epidemiologic data demonstrating a clear association with poor pregnancy outcomes (6, 7). However, histopathological studies of human fetuses and placenta were not performed in these studies. To overcome this limitation, we previously inoculated human second-trimester placenta explants with wild-type RVFV and found that villi were susceptible to infection with RVFV strain ZH501 (31). The villous cytotrophoblasts and syncytiotrophoblast were particularly permissive to RVFV. This finding was alarming because syncytiotrophoblast is generally resistant to most TORCH pathogens (40) by providing a large physical barrier (41 –43) or secreting anti-microbial factors (25, 44). In a separate study, human term villous explants were also permissive to infection with RVFV strain 35/74 (2).

Given the data from our lab and others that RVFV can replicate in human explants, here, we conducted a direct comparison between human and sheep tissues regarding permissivity to infection with RVFV and LAVs. We obtained villi, chorioallantoic, and allantoic membranes from freshly harvested sheep placentomes and dissected them into explants of comparable size. Sheep explant cultures containing villi produced substantial levels of RVFV ZH501 viral RNA over 72 hours—approximately 100-fold higher than that produced by human villi. All three of the human tissues we tested (decidua, fetal membrane, and villi) were also permissive to wild-type RVFV ZH501, although virus production had delayed kinetics and lower overall production compared to ovine counterparts. In our model of ex vivo placental infection, human villi appeared more permissive to ZH501 infection than the decidua or fetal membranes. Overall, sheep explants were more permissive to RVFV infection, and this is reflective of the very high rates of fetal infection in pregnant sheep naturally infected with RVFV. This difference in permissivity and replicative capacity could be due to differences in viral receptor expression and/or innate immune responses and warrants further mechanistic studies.

The high rates of mortality in livestock and morbidity in humans during RVFV outbreaks cause significant socioeconomic strain in affected areas (1). To reduce the impacts of RVF disease in livestock and incidental transmission to humans, LAVs have been developed for use in ruminant livestock. While LAVs are effective at preventing infection after challenge, the safety of some LAVs may be questionable as several have been shown to display residual virulence in pregnant sheep (10, 11, 16, 45). Here, we used the placental explant model to study the permissivity of sheep and human tissues to infection with several relevant LAVs. In sheep placenta explants, LAVs (ΔNSs, MP-12, and ΔNSs/ΔNSm) replicated to moderate levels in villi and fetal membranes, albeit significantly less than wild-type strain ZH501. ΔNSm, on the other hand, generally replicated as well as ZH501 in both sheep and human explants, which is consistent with other in vivo studies showing ΔNSm retains significant neurovirulence (46). Human villous cultures infected with ΔNSs, MP-12, and ΔNSs/ΔNSm undergo early replication, which plateaued by 36 hpi. Consistent with these observations, villous and decidua culture supernatant infected with ΔNSs or ΔNSs/ΔNSm had elevated levels of IFN-α and IFN-λ1 72 hpi compared to uninfected and ZH501-infected controls. ZH501 and ΔNSm, which express NSs, were infectious in the villous, decidua, and fetal membrane cultures, and neither IFN-α nor IFN-λ1 were detected in the supernatant. From these results, we can conclude that the nonstructural protein, NSs, contributes to virulence at the maternal–fetal interface by antagonizing the expression of type I interferons, cytokines that contribute to an anti-viral state within the cell and surrounding areas (47). These results also suggest that NSs inhibits type III interferons in the villous, which is further corroborated by another in vitro study that showed exogenous RVFV NSs inhibits interferon-stimulated response element activation in response to IFN-λ1 (48). It is promising to see human placenta explants control LAV infection; however, if these attenuated viruses are not controlled in the periphery and reach or breach the maternal–fetal interface, the local immune response could contribute to adverse effects on the developing fetus. For instance, the type I and type III interferon responses stimulated by the presence of RVFV might lead to adverse pathological effects as seen with Zika virus infection in mice (32).

Live-attenuated vaccines for human use have not yet been approved, but a few have advanced into clinical trials. In 2006–2008, the attenuated MP-12 strain was included in a phase I/II clinical trial, but it has yet to progress further to the author’s knowledge (14). Other LAV candidates, including the ΔNSs/ΔNSm and a four-segmented vaccine approach (RVFV-4s), have or will soon enter early-stage clinical trials in humans (21, 22). Given the evidence of vertical transmission of RVFV in pregnant women and direct infection of the human placenta in culture, it is important to evaluate the risk of infection and subsequent pathogenesis of human placentas to RVFV LAVs. Furthermore, early-stage vaccine clinical safety trials typically do not include pregnant women due to safety and ethical concerns, making the study of these questions in a laboratory setting even more crucial. Placenta explant cultures are also a tractable model to use in a BSL-3 setting. The use of human and sheep placenta explants, studied at various stages of gestation, could provide pivotal information regarding the cell types targeted by RVFV and the local immune responses to infection. It is important to emphasize that these explant cultures are an incomplete yet biologically relevant model due to the lack of a circulatory system and cross-body communication. Regardless, explants provide a diverse local cellular environment, structure, and intertissue cellular communication that immortalized cell cultures cannot.

RVFV has caused outbreaks with high rates of abortion among sheep and other ruminant livestock for almost a century (37, 38). In contrast, the apparent impacts on pregnant women appear to be significantly lower, but limited epidemiological studies examining pregnancy outcomes in women in endemic regions may account for this perceived difference. We found here that villi from both species are highly susceptible to RVFV infection as evident by the increasing levels of virus over time and the antigen detection. Once RVFV reaches the chorionic villi, it would likely be difficult to control infection and spread to the fetus given the proximity to the fetal endothelium and blood supply (27). Interestingly, the fetal membranes from sheep produced high levels of virus, whereas RVFV did not replicate well in human fetal membrane cultures. Looking closer at one of the human fetal membranes, there is a stark difference in the regions that are infected with RVFV ZH501. The chorionic trophoblasts, which naturally face the mother, were diffusely infected with ZH501, whereas the amniotic epithelium, which faces the fetus, had very little infection. The nature of the explant culture provides equal exposure of all sides of the explant to virus, preventing an infection bias. The fetal membrane in humans, particularly the amniotic epithelium and stroma, which only had a few regions of punctate infection, may play an important role in preventing vertical transmission and protecting the fetus. Surprisingly, fetal membrane cultures did not produce high concentrations of IFN-α or IFN-λ1, suggesting the resistance to infection could be mediated by a different mechanism. Sheep allantoic and chorionic membranes appear to be equally vulnerable to infection, which might explain the higher incidence of fetal demise in livestock compared to humans. In vivo studies using reporter viruses in pregnant sheep and rats could delineate the spatial and temporal aspects of vertical transmission of RVFV. This study is limited by the low number of biological replicates due to the difficulty in procuring sheep and human placentas; thus, additional studies are warranted to corroborate our findings. Furthermore, given multiple studies have shown that virus dose and gestational age may influence disease outcomes (2, 10, 16), future studies should be performed to determine whether these factors could affect vertical transmission across the placenta. Further explant studies should also be performed to understand what makes human amniotic epithelial cells more resistant to RVFV in order to develop targeted therapeutics to prevent or minimize the pathogenic effects following vertical transmission.

In summary, we have used sheep and human placenta explant cultures as a tool to identify cells and tissues targeted by RVFV at the maternal–fetal interface. We provide evidence that although LAVs can infect the decidua, fetal membrane, and villi of humans, spatial restraints, particularly the decidua and fetal membrane, could play an important role in controlling infection and preventing virus dissemination to the fetus.

MATERIALS AND METHODS

Viruses

Virulent RVFV strain ZH501 was generated from reverse genetics plasmids. The recovered virus was provided to our laboratory by B. Miller (CDC, Ft. Collins, CO) and S. Nichol (CDC, Atlanta, GA). RVFV strains with gene deletions (ΔNSs, ΔNSm, and ΔNSs/ΔNSm) were provided by A.K. McElroy (University of Pittsburgh). MP-12 was generously provided by Ted Ross (Cleveland Clinic Florida Research & Innovation Center, Port St. Lucie, FL). All strains of RVFV were propagated on Vero E6 cells following standard methods, and the stock titer was determined by a standard viral plaque assay explained in McMillen et al. (31). For explant studies, stock virus was diluted in virus growth medium [Dulbecco’s modified Eagle medium (DMEM), 2% (vol/vol) fetal bovine serum (FBS), 1% L-glutamine, and 1% penicillin–streptomycin] to the desired dose. Uninfected controls were mock-inoculated with virus growth medium.

Biosafety

Research with RVFV strains ZH501, ΔNSs, and ΔNSm was performed at biosafety level 3 (BSL-3) in the University of Pittsburgh Regional Biocontainment Laboratory (RBL). The University of Pittsburgh RBL is registered with the Centers for Disease Control and Prevention and the United States Department of Agriculture for work with select agents. Work with RVFV strains MP-12 and ΔNSs/ΔNSm, which are both excluded from the DSAT Select Agent lists, was performed at biosafety level 2 (BSL-2) as per local Institutional Biosafety Committee guidelines.

Placenta procurement

Sheep (Texel crossed with Dorset or Suffolk breeds) placentas were collected by George and Lisa Wherry at Wherry Farms, Scenery Hill, PA. Placentas were collected from the field or barn or extracted directly from the sheep. Most of the sheep placentas were procured at term (n = 3; approximately 5 months gestation) after a natural delivery, whereas one placenta was obtained from a pre-term delivery (4 months gestation). Histopathological analysis of the pre-term sheep placenta did not show signs of lesions. Immediately after collection, placentas were placed in complete growth media consisting of DMEM/F12 (1:1), 10% FBS, 2% penicillin–streptomycin, and 250 µg/mL of amphotericin B to minimize the growth of microbial contaminants.

Explant studies

All placentas were dissected within 6–12 hours of procurement, and dissection was conducted based on tissue type [humans: decidua, villi, and fetal membranes (chorion and/or amnion); sheep: chorionic membrane, allantoic membrane, and villi]. Placenta tissues, approximately 5 × 5 mm sections, were placed in 24-well plates, covered in complete growth medium, and allowed to rest overnight at 37°C, 5% CO₂. On the day of infection, the complete growth media were removed from each placenta section prior to inoculation with 200 µL of RVFV (1 × 10⁵ pfu). Placenta sections were incubated for 1 hour at 37°C, 5% CO₂ to allow for virus adsorption. Following the adsorption period, the inoculum was removed, tissue was washed once with 1× PBS, and then, 1 mL of virus growth medium was added to all samples. The plate was cultured for 48 or 72 hours, and 100 µL of culture supernatant was pooled from one to two wells from each tissue daily at 0, 24, 36, 48, and 72 hpi (31). For each human placenta donor (n = 3), one to two samples were analyzed from pooled culture supernatant at each time point. For each sheep placenta donor (n = 4), three samples were analyzed from pooled culture supernatant at each time point.

RNA isolation and q-RT-PCR

The culture supernatant (50 µL) was inactivated in 900 µL of TRIzol reagent (Invitrogen). RNA isolation and q-RT-PCR were performed as described (31).

Enzyme-linked immunosorbent assay

Supernatant collected from human tissues at 48 or 72 hpi after infection with RVFV ZH501 or LAVs (ΔNSs, ΔNSm, MP-12, and ΔNSs/ΔNSm) was used to detect total human interferon alpha (IFN-α; Invitrogen catalog # BMS216) and human interleukin 29 (IL-29/IFN-λ1; Abcam # ab100568) following the manufacturer’s instructions. Standard curves were generated to quantitate protein concentrations. The LOD for the IFN-α and IFN-λ1 assays was 3.2 and 13.72 pg/mL, respectively. For each human placenta donor (n = 3), one to two samples were analyzed from pooled culture supernatant at either time point.

Immunohistochemistry and microscopy

Human and sheep tissues were fixed in 4% paraformaldehyde (PFA) for 24 hours then paraffin-embedded by standard protocols. Tissue sections 6 µm thick were embedded on slides and baked overnight at 65°C. Slides were deparaffinized using a standard xylene and alcohol rehydration series, then boiled in 10 mM citric acid buffer (pH 6.0) to unmask antigen-binding epitopes. Tissue sections were then permeabilized using 0.1% Triton X-100 detergent in phosphate buffered saline (PBS). Tissue sections were blocked in Bloxall blocking solution (Vector Laboratories) for 10 minutes, then washed 3× in PBS. Then, using the Vectastain Elite ABC Kit, Peroxidase (Rabbit IgG) and Vector NovaRED Substrate Kit, Peroxidase (Vector Laboratories), tissue sections were blocked in normal blocking serum, avidin block, and biotin block for 15–20 minutes, with five washes in 1× PBS between each block. Following five washes in PBS, anti-RVFV nucleoprotein rabbit IgG (1:100 dilution; custom-made via Genscript) was applied to each sample and incubated for 1 hour. Tissues were washed five times in PBS; then, the Vector Biotinylated Goat anti-Rabbit secondary antibody was applied for 30 minutes. Tissues were washed five times with PBS; then, the Vectastain ABC-HRP reagent was added for 30 minutes. The tissues were washed five times; then, the Vector NovaRED was applied for 15 minutes. The Vector NovaRED was removed by dunking slides into fresh water. Tissues were counterstained in 50% diluted hematoxylin (Leica, catalog # 3801575) for 20 seconds, washed in water, dunked in 5% acetic acid five times, washed in water, and then incubated in Scott’s tap water (Cancer Diagnostics Inc., catalog # CM4951W) for 10 minutes to promote counterstain bluing. After dehydrating in Clear-Rite (Thermo Scientific, catalog #6901), coverslips were mounted over the tissues using toluene. Slides dried overnight before imaging. All steps were performed at room temperature unless stated otherwise. Primary delete and no infection controls were used to identify nonspecific binding of the secondary detection antibody or all reagents, respectively. Red immunopositivity observed above the no infection control was deemed as positive for RVFV antigen. Hematoxylin staining provided a structural stain to identify immune cells. IHC slides were imaged by light (optical) microscopy using a Nikon NI-E confocal microscope at the University of Pittsburgh Center for Biological Imaging or the Olympus LCmicro Software at the One Health Institute at the University of California Davis. Denoising and contrasting were performed using Adobe Photoshop. Tissue samples were blinded and analyzed by a veterinary pathologist or a licensed pathologist specializing in placental pathology.

Statistics

For virus replication curves and cytokine expression assays, one-way or two-way ANOVA with multiple comparisons was performed using GraphPad Prism (version 8.0). To calculate virus replication rates, simple linear regression from log(10) transformed data was performed starting at 24 or 36 hpi for sheep and human cultures, respectively.

ACKNOWLEDGMENTS

The authors would like to thank the Center for Biological Imaging, particularly Michael J Calderon and Katherine E. Helfrich, for assistance with image acquisition. We recognize the McGowan Center for Regenerative Medicine for histology support. Hayley Nordstrom provided exceptional original artwork. This work would not have been possible without the generous help from George and Lisa Wherry at Wherry Farms, Scenery Hill, PA.

All work was supported by the National Institutes of Health (R01 AI150792 and R01 AI150792S1 to A.L.H., T32 AI060525 and K01 AI165965 to C.M.M). NIH award (UC7AI180311) from the National Institute of Allergy and Infectious Diseases (NIAID) supported the Operations of the University of Pittsburgh Regional Biocontainment Laboratory (RBL) within the Center for Vaccine Research (CVR). The funders had no role in the study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Conceptualization was performed by Cynthia M. McMillen and Amy L. Hartman; data curation was performed by Cynthia M. McMillen, Ryan M. Hoehl, Devin A Boyles, Jackson J. McGaughey, Lauren Skvarca, and Amy L. Hartman; formal analysis was performed by Cynthia M. McMillen, Christina Megli, Rebecca Radisic, Lauren B. Skvarca, Brian H Bird, and Amy L. Hartman; funding acquisition was performed by Amy L. Hartman; investigation was performed by Cynthia M. McMillen, Rebecca Radisic, Lauren B. Skvarca, Ryan M. Hoehl, Devin A. Boyles, Jackson J. McGaughey, and Amy L. Hartman; methodology was performed by Cynthia M. McMillen, Christina Megli, Rebecca Radisic, Lauren B. Skvarca, Ryan M. Hoehl, Jackson J. McGaughey, Brian H. Bird, and Anita K McElroy; project administration was performed by Amy L. Hartman; supervision was performed by Amy L. Hartman; validation was performed by Amy L. Hartman; visualization was performed by Cynthia M. McMillen, Rebecca Radisic, Lauren B. Skvarca, Brian H. Bird, and Amy L. Hartman; writing (original draft) was performed by Cynthia M. McMillen and Amy L. Hartman.

Contributor Information

Cynthia M. McMillen, Email: cmm287@pitt.edu.

Amy L. Hartman, Email: hartman2@pitt.edu.

Stacey Schultz-Cherry, St. Jude Children's Research Hospital, Memphis, Tennessee, USA.

ETHICS APPROVAL

Human placentas isolated between 32 and 36 weeks gestation via cesarean section were obtained from the Steven C Caritis Magee Obstetric Maternal and Infant Biobank through an honest broker system within 30 minutes of delivery. Tissue procurement was approved by the University of Pittsburgh Institutional Review Board (IRB) 19100322.

DATA AVAILABILITY

All data needed to evaluate the conclusions in the paper are present in the paper and/or the supplemental material. Additional data related to this paper may be requested from the authors.

SUPPLEMENTAL MATERIAL

The following material is available online at https://doi.org/10.1128/jvi.00983-24.

Supplemental figures and tables. jvi.00983-24-s0001.pdf.

Fig. S1-S3; Tables S1 and S2.

jvi.00983-24-s0001.pdf^{(2MB, pdf)}

DOI: 10.1128/jvi.00983-24.SuF1

ASM does not own the copyrights to Supplemental Material that may be linked to, or accessed through, an article. The authors have granted ASM a non-exclusive, world-wide license to publish the Supplemental Material files. Please contact the corresponding author directly for reuse.

REFERENCES

1. Chengula AA, Mdegela RH, Kasanga CJ. 2013. Socio-economic impact of Rift Valley fever to pastoralists and agro pastoralists in Arusha, Manyara and Morogoro regions in Tanzania. Springerplus 2:549. doi: 10.1186/2193-1801-2-549 [DOI] [PMC free article] [PubMed] [Google Scholar]
2. Oymans J, Wichgers Schreur PJ, van Keulen L, Kant J, Kortekaas J. 2020. Rift Valley fever virus targets the maternal-foetal interface in ovine and human placentas. PLoS Negl Trop Dis 14:e0007898. doi: 10.1371/journal.pntd.0007898 [DOI] [PMC free article] [PubMed] [Google Scholar]
3. Coetzer JA. 1977. The pathology of Rift Valley fever. I. Lesions occurring in natural cases in new-born lambs. Onderstepoort J Vet Res 44:205–211. [PubMed] [Google Scholar]
4. Coetzer JA. 1982. The pathology of Rift Valley fever. II. Lesions occurring in field cases in adult cattle, calves and aborted foetuses. Onderstepoort J Vet Res 49:11–17. [PubMed] [Google Scholar]
5. Baudin M, Jumaa AM, Jomma HJE, Karsany MS, Bucht G, Näslund J, Ahlm C, Evander M, Mohamed N. 2016. Association of Rift Valley fever virus infection with miscarriage in sudanese women: a cross-sectional study. Lancet Glob Health 4:e864–e871. doi: 10.1016/S2214-109X(16)30176-0 [DOI] [PubMed] [Google Scholar]
6. Adam I, Karsany MS. 2008. Case report: Rift Valley fever with vertical transmission in a pregnant sudanese woman. J Med Virol 80:929. doi: 10.1002/jmv.21132 [DOI] [PubMed] [Google Scholar]
7. Arishi HM, Aqeel AY, Al Hazmi MM. 2006. Vertical transmission of fatal Rift Valley fever in a newborn. Ann Trop Paediatr 26:251–253. doi: 10.1179/146532806X120363 [DOI] [PubMed] [Google Scholar]
8. Smithburn KC. 1949. Rift Valley fever; the neurotropic adaptation of the virus and the experimental use of this modified virus as a vaccine. Br J Exp Pathol 30:1–16. [PMC free article] [PubMed] [Google Scholar]
9. Muller R, Saluzzo JF, Lopez N, Dreier T, Turell M, Smith J, Bouloy M. 1995. Characterization of clone 13, a naturally attenuated avirulent isolate of Rift Valley fever virus, which is altered in the small segment. Am J Trop Med Hyg 53:405–411. doi: 10.4269/ajtmh.1995.53.405 [DOI] [PubMed] [Google Scholar]
10. Makoschey B, van Kilsdonk E, Hubers WR, Vrijenhoek MP, Smit M, Wichgers Schreur PJ, Kortekaas J, Moulin V. 2016. Rift Valley fever vaccine virus clone 13 is able to cross the ovine placental barrier associated with foetal infections, malformations, and stillbirths. PLoS Negl Trop Dis 10:e0004550. doi: 10.1371/journal.pntd.0004550 [DOI] [PMC free article] [PubMed] [Google Scholar]
11. Botros B, Omar A, Elian K, Mohamed G, Soliman A, Salib A, Salman D, Saad M, Earhart K. 2006. Adverse response of non-indigenous cattle of European breeds to live attenuated Smithburn Rift Valley fever vaccine. J Med Virol 78:787–791. doi: 10.1002/jmv.20624 [DOI] [PubMed] [Google Scholar]
12. Lokugamage N, Ikegami T. 2017. Genetic stability of Rft Valley fever virus MP-12 vaccine during serial passages in culture cells. NPJ Vaccines 2:20. doi: 10.1038/s41541-017-0021-9 [DOI] [PMC free article] [PubMed] [Google Scholar]
13. Ikegami T, Hill TE, Smith JK, Zhang L, Juelich TL, Gong B, Slack OAL, Ly HJ, Lokugamage N, Freiberg AN. 2015. Rift Valley fever virus MP-12 vaccine is fully attenuated by a combination of partial attenuations in the S, M, and L segments. J Virol 89:7262–7276. doi: 10.1128/JVI.00135-15 [DOI] [PMC free article] [PubMed] [Google Scholar]
14. Pittman PR, Norris SL, Brown ES, Ranadive MV, Schibly BA, Bettinger GE, Lokugamage N, Korman L, Morrill JC, Peters CJ. 2016. Rift Valley fever MP-12 vaccine phase 2 clinical trial: safety, immunogenicity, and genetic characterization of virus isolates. Vaccine 34:523–530. doi: 10.1016/j.vaccine.2015.11.078 [DOI] [PMC free article] [PubMed] [Google Scholar]
15. Pittman PR, McClain D, Quinn X, Coonan KM, Mangiafico J, Makuch RS, Morrill J, Peters CJ. 2016. Safety and immunogenicity of a mutagenized, live attenuated Rift Valley fever vaccine, MP-12, in a phase 1 dose escalation and route comparison study in humans. Vaccine 34:424–429. doi: 10.1016/j.vaccine.2015.12.030 [DOI] [PubMed] [Google Scholar]
16. Hunter P, Erasmus BJ, Vorster JH. 2002. Teratogenicity of a mutagenised Rift Valley fever virus (MVP 12) in sheep. Onderstepoort J Vet Res 69:95–98. [PubMed] [Google Scholar]
17. Morrill JC, Carpenter L, Taylor D, Ramsburg HH, Quance J, Peters CJ. 1991. Further evaluation of a mutagen-attenuated Rift Valley fever vaccine in sheep. Vaccine 9:35–41. doi: 10.1016/0264-410x(91)90314-v [DOI] [PubMed] [Google Scholar]
18. Morrill JC, Jennings GB, Caplen H, Turell MJ, Johnson AJ, Peters CJ. 1987. Pathogenicity and immunogenicity of a mutagen-attenuated Rift Valley fever virus immunogen in pregnant ewes. Am J Vet Res 48:1042–1047. [PubMed] [Google Scholar]
19. Bird BH, Maartens LH, Campbell S, Erasmus BJ, Erickson BR, Dodd KA, Spiropoulou CF, Cannon D, Drew CP, Knust B, McElroy AK, Khristova ML, Albariño CG, Nichol ST. 2011. Rift Valley fever virus vaccine lacking the Nss and NSM genes is safe, nonteratogenic, and confers protection from viremia, pyrexia, and abortion following challenge in adult and pregnant sheep. J Virol 85:12901–12909. doi: 10.1128/JVI.06046-11 [DOI] [PMC free article] [PubMed] [Google Scholar]
20. Jenkin D, Wright D, Folegatti PM, Platt A, Poulton I, Lawrie A, Tran N, Boyd A, Turner C, Gitonga JN, Karanja HK, Mugo D, Ewer KJ, Bowden TA, Gilbert SC, Charleston B, Kaleebu P, Hill AVS, Warimwe GM. 2023. Safety and immunogenicity of a chadox1 vaccine against Rift Valley fever in UK adults: an open-label, non-randomised, first-in-human phase 1 clinical trial. Lancet Infect Dis 23:956–964. doi: 10.1016/S1473-3099(23)00068-3 [DOI] [PMC free article] [PubMed] [Google Scholar]
21. Rogers J. 2022. Rift Valley fever vaccines to advance with new $50 million CEPI and EU funding call, on coalition for epidemic preparedness innovations. Available from: https://cepi.net/news_cepi/rift-valley-fever-vaccines-to-advance-with-new-50-million-cepi-and-eu-funding-call. Retrieved 27 Jan 2023.
22. Rogers J. 2023. CEPI partners with university of California, Davis to advance a vaccine against potentially deadly rift valley fever virus into clinical trials, on coalition for epidemic preparedness innovations. Available from: https://cepi.net/news_cepi/cepi-partners-with-university-of-california-davis-to-advance-a-vaccine-against-potentially-deadly-rift-valley-fever-virus-into-clinical-trials. Retrieved 5 Jan 2024.
23. Pijnenborg R, Robertson WB, Brosens I, Dixon G. 1981. Review article: trophoblast invasion and the establishment of haemochorial placentation in man and laboratory animals. Placenta 2:71–91. doi: 10.1016/s0143-4004(81)80042-2 [DOI] [PubMed] [Google Scholar]
24. Wooding P, Burton G. 2008. Comparative placentation. 1 ed. Springer, Berlin, Heidelberg. [Google Scholar]
25. Corry J, Arora N, Good CA, Sadovsky Y, Coyne CB. 2017. Organotypic models of type III interferon-mediated protection from Zika virus infections at the maternal-fetal interface. Proc Natl Acad Sci U S A 114:9433–9438. doi: 10.1073/pnas.1707513114 [DOI] [PMC free article] [PubMed] [Google Scholar]
26. Fahmi A, Brügger M, Démoulins T, Zumkehr B, Oliveira Esteves BI, Bracher L, Wotzkow C, Blank F, Thiel V, Baud D, Alves MP. 2021. SARS-CoV-2 can infect and propagate in human placenta explants. Cell Rep Med 2:100456. doi: 10.1016/j.xcrm.2021.100456 [DOI] [PMC free article] [PubMed] [Google Scholar]
27. Leiser R, Kaufmann P. 1994. Placental structure: in a comparative aspect. Exp Clin Endocrinol 102:122–134. doi: 10.1055/s-0029-1211275 [DOI] [PubMed] [Google Scholar]
28. Medina L, Guerrero-Muñoz J, Castillo C, Liempi A, Fernández-Moya A, Araneda S, Ortega Y, Rivas C, Maya JD, Kemmerling U. 2022. Differential microRNAs expression during ex vivo infection of canine and ovine placental explants with Trypanosoma cruzi and Toxoplasma gondii. Acta Trop 235:106651. doi: 10.1016/j.actatropica.2022.106651 [DOI] [PubMed] [Google Scholar]
29. Liempi A, Castillo C, Medina L, Galanti N, Maya JD, Parraguez VH, Kemmerling U. 2020. Comparative ex vivo infection with Trypanosoma cruzi and Toxoplasma gondii of human, canine and ovine placenta: analysis of tissue damage and infection efficiency. Parasitol Int 76:102065. doi: 10.1016/j.parint.2020.102065 [DOI] [PubMed] [Google Scholar]
30. Horcajo P, Ortega-Mora LM, Benavides J, Sánchez-Sánchez R, Amieva R, Collantes-Fernández E, Pastor-Fernández I. 2023. Ovine placental explants: a new ex vivo model to study host‒pathogen interactions in reproductive pathogens. Theriogenology 212:157–171. doi: 10.1016/j.theriogenology.2023.09.009 [DOI] [PubMed] [Google Scholar]
31. McMillen CM, Arora N, Boyles DA, Albe JR, Kujawa MR, Bonadio JF, Coyne CB, Hartman AL. 2018. Rift Valley fever virus induces fetal demise in Sprague-Dawley rats through direct placental infection. Sci Adv 4:eaau9812. doi: 10.1126/sciadv.aau9812 [DOI] [PMC free article] [PubMed] [Google Scholar]
32. Casazza RL, Lazear HM, Miner JJ. 2020. Protective and pathogenic effects of interferon signaling during pregnancy. Viral Immunol 33:3–11. doi: 10.1089/vim.2019.0076 [DOI] [PMC free article] [PubMed] [Google Scholar]
33. Megli CJ, Coyne CB. 2022. Infections at the maternal-fetal interface: an overview of pathogenesis and defence. Nat Rev Microbiol 20:67–82. doi: 10.1038/s41579-021-00610-y [DOI] [PMC free article] [PubMed] [Google Scholar]
34. Wells AI, Coyne CB. 2018. Type III Interferons in antiviral defenses at barrier surfaces. Trends Immunol 39:848–858. doi: 10.1016/j.it.2018.08.008 [DOI] [PMC free article] [PubMed] [Google Scholar]
35. Ding J, Maxwell A, Adzibolosu N, Hu A, You Y, Liao A, Mor G. 2022. Mechanisms of immune regulation by the placenta: role of type I interferon and interferon-stimulated genes signaling during pregnancy. Immunol Rev 308:9–24. doi: 10.1111/imr.13077 [DOI] [PMC free article] [PubMed] [Google Scholar]
36. Koi H, Zhang J, Parry S. 2001. The mechanisms of placental viral infection. Ann N Y Acad Sci 943:148–156. doi: 10.1111/j.1749-6632.2001.tb03798.x [DOI] [PubMed] [Google Scholar]
37. Daubney R, Hudson JR, Garnham PC. 1931. Enzootic hepatitis or rift valley fever. an undescribted virus disease in sheep cattle and man from East Africa. Pathol 34:545–579. doi: 10.1002/path.1700340418 [DOI] [Google Scholar]
38. Findlay GM, Daubney R. 1931. The virus of Rift Valley fever or enzootic hepatitis. Lancet 218:1350–1351. doi: 10.1016/S0140-6736(00)99624-7 [DOI] [Google Scholar]
39. Odendaal L, Clift SJ, Fosgate GT, Davis AS. 2020. Ovine fetal and placental lesions and cellular tropism in natural Rift Valley fever virus infections. Vet Pathol 57:791–806. doi: 10.1177/0300985820954549 [DOI] [PubMed] [Google Scholar]
40. Robbins JR, Skrzypczynska KM, Zeldovich VB, Kapidzic M, Bakardjiev AI. 2010. Placental syncytiotrophoblast constitutes a major barrier to vertical transmission of Listeria monocytogenes. PLoS Pathog 6:e1000732. doi: 10.1371/journal.ppat.1000732 [DOI] [PMC free article] [PubMed] [Google Scholar]
41. Koi H, Zhang J, Makrigiannakis A, Getsios S, MacCalman CD, Strauss JF, Parry S. 2002. Syncytiotrophoblast is a barrier to maternal-fetal transmission of herpes simplex virus. Biol Reprod 67:1572–1579. doi: 10.1095/biolreprod.102.004325 [DOI] [PubMed] [Google Scholar]
42. Zeldovich VB, Clausen CH, Bradford E, Fletcher DA, Maltepe E, Robbins JR, Bakardjiev AI. 2013. Placental syncytium forms a biophysical barrier against pathogen invasion. PLoS Pathog 9:e1003821. doi: 10.1371/journal.ppat.1003821 [DOI] [PMC free article] [PubMed] [Google Scholar]
43. Ander SE, Rudzki EN, Arora N, Sadovsky Y, Coyne CB, Boyle JP. 2018. Human placental syncytiotrophoblasts restrict Toxoplasma gondii attachment and replication and respond to infection by producing immunomodulatory chemokines. mBio 9:e01678-17. doi: 10.1128/mBio.01678-17 [DOI] [PMC free article] [PubMed] [Google Scholar]
44. Bayer A, Lennemann NJ, Ouyang Y, Bramley JC, Morosky S, Marques ETDA Jr, Cherry S, Sadovsky Y, Coyne CB. 2016. Type III interferons produced by human placental trophoblasts confer protection against Zika virus infection. Cell Host Microbe 19:705–712. doi: 10.1016/j.chom.2016.03.008 [DOI] [PMC free article] [PubMed] [Google Scholar]
45. Coetzer JA, Barnard BJ. 1977. Hydrops amnii in sheep associated with hydranencephaly and arthrogryposis with wesselsbron disease and rift valley fever viruses as aetiological agents. Onderstepoort J Vet Res 44:119–126. [PubMed] [Google Scholar]
46. Bird BH, Albariño CG, Nichol ST. 2007. Rift Valley fever virus lacking NSM proteins retains high virulence in vivo and may provide a model of human delayed onset neurologic disease. Virology 362:10–15. doi: 10.1016/j.virol.2007.01.046 [DOI] [PubMed] [Google Scholar]
47. Billecocq A, Spiegel M, Vialat P, Kohl A, Weber F, Bouloy M, Haller O. 2004. Nss protein of Rift Valley fever virus blocks interferon production by inhibiting host gene transcription. J Virol 78:9798–9806. doi: 10.1128/JVI.78.18.9798-9806.2004 [DOI] [PMC free article] [PubMed] [Google Scholar]
48. Koenig ZT. 2018. Type III interferon control of Rift Valley fever virus at epithelial cell barrier. Master of Science. University of Pittsburgh, SUNY Geneseo. [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplemental figures and tables. jvi.00983-24-s0001.pdf.

Fig. S1-S3; Tables S1 and S2.

jvi.00983-24-s0001.pdf^{(2MB, pdf)}

DOI: 10.1128/jvi.00983-24.SuF1

Data Availability Statement

All data needed to evaluate the conclusions in the paper are present in the paper and/or the supplemental material. Additional data related to this paper may be requested from the authors.

[B1] 1. Chengula AA, Mdegela RH, Kasanga CJ. 2013. Socio-economic impact of Rift Valley fever to pastoralists and agro pastoralists in Arusha, Manyara and Morogoro regions in Tanzania. Springerplus 2:549. doi: 10.1186/2193-1801-2-549 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B2] 2. Oymans J, Wichgers Schreur PJ, van Keulen L, Kant J, Kortekaas J. 2020. Rift Valley fever virus targets the maternal-foetal interface in ovine and human placentas. PLoS Negl Trop Dis 14:e0007898. doi: 10.1371/journal.pntd.0007898 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B3] 3. Coetzer JA. 1977. The pathology of Rift Valley fever. I. Lesions occurring in natural cases in new-born lambs. Onderstepoort J Vet Res 44:205–211. [PubMed] [Google Scholar]

[B4] 4. Coetzer JA. 1982. The pathology of Rift Valley fever. II. Lesions occurring in field cases in adult cattle, calves and aborted foetuses. Onderstepoort J Vet Res 49:11–17. [PubMed] [Google Scholar]

[B5] 5. Baudin M, Jumaa AM, Jomma HJE, Karsany MS, Bucht G, Näslund J, Ahlm C, Evander M, Mohamed N. 2016. Association of Rift Valley fever virus infection with miscarriage in sudanese women: a cross-sectional study. Lancet Glob Health 4:e864–e871. doi: 10.1016/S2214-109X(16)30176-0 [DOI] [PubMed] [Google Scholar]

[B6] 6. Adam I, Karsany MS. 2008. Case report: Rift Valley fever with vertical transmission in a pregnant sudanese woman. J Med Virol 80:929. doi: 10.1002/jmv.21132 [DOI] [PubMed] [Google Scholar]

[B7] 7. Arishi HM, Aqeel AY, Al Hazmi MM. 2006. Vertical transmission of fatal Rift Valley fever in a newborn. Ann Trop Paediatr 26:251–253. doi: 10.1179/146532806X120363 [DOI] [PubMed] [Google Scholar]

[B8] 8. Smithburn KC. 1949. Rift Valley fever; the neurotropic adaptation of the virus and the experimental use of this modified virus as a vaccine. Br J Exp Pathol 30:1–16. [PMC free article] [PubMed] [Google Scholar]

[B9] 9. Muller R, Saluzzo JF, Lopez N, Dreier T, Turell M, Smith J, Bouloy M. 1995. Characterization of clone 13, a naturally attenuated avirulent isolate of Rift Valley fever virus, which is altered in the small segment. Am J Trop Med Hyg 53:405–411. doi: 10.4269/ajtmh.1995.53.405 [DOI] [PubMed] [Google Scholar]

[B10] 10. Makoschey B, van Kilsdonk E, Hubers WR, Vrijenhoek MP, Smit M, Wichgers Schreur PJ, Kortekaas J, Moulin V. 2016. Rift Valley fever vaccine virus clone 13 is able to cross the ovine placental barrier associated with foetal infections, malformations, and stillbirths. PLoS Negl Trop Dis 10:e0004550. doi: 10.1371/journal.pntd.0004550 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B11] 11. Botros B, Omar A, Elian K, Mohamed G, Soliman A, Salib A, Salman D, Saad M, Earhart K. 2006. Adverse response of non-indigenous cattle of European breeds to live attenuated Smithburn Rift Valley fever vaccine. J Med Virol 78:787–791. doi: 10.1002/jmv.20624 [DOI] [PubMed] [Google Scholar]

[B12] 12. Lokugamage N, Ikegami T. 2017. Genetic stability of Rft Valley fever virus MP-12 vaccine during serial passages in culture cells. NPJ Vaccines 2:20. doi: 10.1038/s41541-017-0021-9 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B13] 13. Ikegami T, Hill TE, Smith JK, Zhang L, Juelich TL, Gong B, Slack OAL, Ly HJ, Lokugamage N, Freiberg AN. 2015. Rift Valley fever virus MP-12 vaccine is fully attenuated by a combination of partial attenuations in the S, M, and L segments. J Virol 89:7262–7276. doi: 10.1128/JVI.00135-15 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B14] 14. Pittman PR, Norris SL, Brown ES, Ranadive MV, Schibly BA, Bettinger GE, Lokugamage N, Korman L, Morrill JC, Peters CJ. 2016. Rift Valley fever MP-12 vaccine phase 2 clinical trial: safety, immunogenicity, and genetic characterization of virus isolates. Vaccine 34:523–530. doi: 10.1016/j.vaccine.2015.11.078 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B15] 15. Pittman PR, McClain D, Quinn X, Coonan KM, Mangiafico J, Makuch RS, Morrill J, Peters CJ. 2016. Safety and immunogenicity of a mutagenized, live attenuated Rift Valley fever vaccine, MP-12, in a phase 1 dose escalation and route comparison study in humans. Vaccine 34:424–429. doi: 10.1016/j.vaccine.2015.12.030 [DOI] [PubMed] [Google Scholar]

[B16] 16. Hunter P, Erasmus BJ, Vorster JH. 2002. Teratogenicity of a mutagenised Rift Valley fever virus (MVP 12) in sheep. Onderstepoort J Vet Res 69:95–98. [PubMed] [Google Scholar]

[B17] 17. Morrill JC, Carpenter L, Taylor D, Ramsburg HH, Quance J, Peters CJ. 1991. Further evaluation of a mutagen-attenuated Rift Valley fever vaccine in sheep. Vaccine 9:35–41. doi: 10.1016/0264-410x(91)90314-v [DOI] [PubMed] [Google Scholar]

[B18] 18. Morrill JC, Jennings GB, Caplen H, Turell MJ, Johnson AJ, Peters CJ. 1987. Pathogenicity and immunogenicity of a mutagen-attenuated Rift Valley fever virus immunogen in pregnant ewes. Am J Vet Res 48:1042–1047. [PubMed] [Google Scholar]

[B19] 19. Bird BH, Maartens LH, Campbell S, Erasmus BJ, Erickson BR, Dodd KA, Spiropoulou CF, Cannon D, Drew CP, Knust B, McElroy AK, Khristova ML, Albariño CG, Nichol ST. 2011. Rift Valley fever virus vaccine lacking the Nss and NSM genes is safe, nonteratogenic, and confers protection from viremia, pyrexia, and abortion following challenge in adult and pregnant sheep. J Virol 85:12901–12909. doi: 10.1128/JVI.06046-11 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B20] 20. Jenkin D, Wright D, Folegatti PM, Platt A, Poulton I, Lawrie A, Tran N, Boyd A, Turner C, Gitonga JN, Karanja HK, Mugo D, Ewer KJ, Bowden TA, Gilbert SC, Charleston B, Kaleebu P, Hill AVS, Warimwe GM. 2023. Safety and immunogenicity of a chadox1 vaccine against Rift Valley fever in UK adults: an open-label, non-randomised, first-in-human phase 1 clinical trial. Lancet Infect Dis 23:956–964. doi: 10.1016/S1473-3099(23)00068-3 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B21] 21. Rogers J. 2022. Rift Valley fever vaccines to advance with new $50 million CEPI and EU funding call, on coalition for epidemic preparedness innovations. Available from: https://cepi.net/news_cepi/rift-valley-fever-vaccines-to-advance-with-new-50-million-cepi-and-eu-funding-call. Retrieved 27 Jan 2023.

[B22] 22. Rogers J. 2023. CEPI partners with university of California, Davis to advance a vaccine against potentially deadly rift valley fever virus into clinical trials, on coalition for epidemic preparedness innovations. Available from: https://cepi.net/news_cepi/cepi-partners-with-university-of-california-davis-to-advance-a-vaccine-against-potentially-deadly-rift-valley-fever-virus-into-clinical-trials. Retrieved 5 Jan 2024.

[B23] 23. Pijnenborg R, Robertson WB, Brosens I, Dixon G. 1981. Review article: trophoblast invasion and the establishment of haemochorial placentation in man and laboratory animals. Placenta 2:71–91. doi: 10.1016/s0143-4004(81)80042-2 [DOI] [PubMed] [Google Scholar]

[B24] 24. Wooding P, Burton G. 2008. Comparative placentation. 1 ed. Springer, Berlin, Heidelberg. [Google Scholar]

[B25] 25. Corry J, Arora N, Good CA, Sadovsky Y, Coyne CB. 2017. Organotypic models of type III interferon-mediated protection from Zika virus infections at the maternal-fetal interface. Proc Natl Acad Sci U S A 114:9433–9438. doi: 10.1073/pnas.1707513114 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B26] 26. Fahmi A, Brügger M, Démoulins T, Zumkehr B, Oliveira Esteves BI, Bracher L, Wotzkow C, Blank F, Thiel V, Baud D, Alves MP. 2021. SARS-CoV-2 can infect and propagate in human placenta explants. Cell Rep Med 2:100456. doi: 10.1016/j.xcrm.2021.100456 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B27] 27. Leiser R, Kaufmann P. 1994. Placental structure: in a comparative aspect. Exp Clin Endocrinol 102:122–134. doi: 10.1055/s-0029-1211275 [DOI] [PubMed] [Google Scholar]

[B28] 28. Medina L, Guerrero-Muñoz J, Castillo C, Liempi A, Fernández-Moya A, Araneda S, Ortega Y, Rivas C, Maya JD, Kemmerling U. 2022. Differential microRNAs expression during ex vivo infection of canine and ovine placental explants with Trypanosoma cruzi and Toxoplasma gondii. Acta Trop 235:106651. doi: 10.1016/j.actatropica.2022.106651 [DOI] [PubMed] [Google Scholar]

[B29] 29. Liempi A, Castillo C, Medina L, Galanti N, Maya JD, Parraguez VH, Kemmerling U. 2020. Comparative ex vivo infection with Trypanosoma cruzi and Toxoplasma gondii of human, canine and ovine placenta: analysis of tissue damage and infection efficiency. Parasitol Int 76:102065. doi: 10.1016/j.parint.2020.102065 [DOI] [PubMed] [Google Scholar]

[B30] 30. Horcajo P, Ortega-Mora LM, Benavides J, Sánchez-Sánchez R, Amieva R, Collantes-Fernández E, Pastor-Fernández I. 2023. Ovine placental explants: a new ex vivo model to study host‒pathogen interactions in reproductive pathogens. Theriogenology 212:157–171. doi: 10.1016/j.theriogenology.2023.09.009 [DOI] [PubMed] [Google Scholar]

[B31] 31. McMillen CM, Arora N, Boyles DA, Albe JR, Kujawa MR, Bonadio JF, Coyne CB, Hartman AL. 2018. Rift Valley fever virus induces fetal demise in Sprague-Dawley rats through direct placental infection. Sci Adv 4:eaau9812. doi: 10.1126/sciadv.aau9812 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B32] 32. Casazza RL, Lazear HM, Miner JJ. 2020. Protective and pathogenic effects of interferon signaling during pregnancy. Viral Immunol 33:3–11. doi: 10.1089/vim.2019.0076 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B33] 33. Megli CJ, Coyne CB. 2022. Infections at the maternal-fetal interface: an overview of pathogenesis and defence. Nat Rev Microbiol 20:67–82. doi: 10.1038/s41579-021-00610-y [DOI] [PMC free article] [PubMed] [Google Scholar]

[B34] 34. Wells AI, Coyne CB. 2018. Type III Interferons in antiviral defenses at barrier surfaces. Trends Immunol 39:848–858. doi: 10.1016/j.it.2018.08.008 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B35] 35. Ding J, Maxwell A, Adzibolosu N, Hu A, You Y, Liao A, Mor G. 2022. Mechanisms of immune regulation by the placenta: role of type I interferon and interferon-stimulated genes signaling during pregnancy. Immunol Rev 308:9–24. doi: 10.1111/imr.13077 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B36] 36. Koi H, Zhang J, Parry S. 2001. The mechanisms of placental viral infection. Ann N Y Acad Sci 943:148–156. doi: 10.1111/j.1749-6632.2001.tb03798.x [DOI] [PubMed] [Google Scholar]

[B37] 37. Daubney R, Hudson JR, Garnham PC. 1931. Enzootic hepatitis or rift valley fever. an undescribted virus disease in sheep cattle and man from East Africa. Pathol 34:545–579. doi: 10.1002/path.1700340418 [DOI] [Google Scholar]

[B38] 38. Findlay GM, Daubney R. 1931. The virus of Rift Valley fever or enzootic hepatitis. Lancet 218:1350–1351. doi: 10.1016/S0140-6736(00)99624-7 [DOI] [Google Scholar]

[B39] 39. Odendaal L, Clift SJ, Fosgate GT, Davis AS. 2020. Ovine fetal and placental lesions and cellular tropism in natural Rift Valley fever virus infections. Vet Pathol 57:791–806. doi: 10.1177/0300985820954549 [DOI] [PubMed] [Google Scholar]

[B40] 40. Robbins JR, Skrzypczynska KM, Zeldovich VB, Kapidzic M, Bakardjiev AI. 2010. Placental syncytiotrophoblast constitutes a major barrier to vertical transmission of Listeria monocytogenes. PLoS Pathog 6:e1000732. doi: 10.1371/journal.ppat.1000732 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B41] 41. Koi H, Zhang J, Makrigiannakis A, Getsios S, MacCalman CD, Strauss JF, Parry S. 2002. Syncytiotrophoblast is a barrier to maternal-fetal transmission of herpes simplex virus. Biol Reprod 67:1572–1579. doi: 10.1095/biolreprod.102.004325 [DOI] [PubMed] [Google Scholar]

[B42] 42. Zeldovich VB, Clausen CH, Bradford E, Fletcher DA, Maltepe E, Robbins JR, Bakardjiev AI. 2013. Placental syncytium forms a biophysical barrier against pathogen invasion. PLoS Pathog 9:e1003821. doi: 10.1371/journal.ppat.1003821 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B43] 43. Ander SE, Rudzki EN, Arora N, Sadovsky Y, Coyne CB, Boyle JP. 2018. Human placental syncytiotrophoblasts restrict Toxoplasma gondii attachment and replication and respond to infection by producing immunomodulatory chemokines. mBio 9:e01678-17. doi: 10.1128/mBio.01678-17 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B44] 44. Bayer A, Lennemann NJ, Ouyang Y, Bramley JC, Morosky S, Marques ETDA Jr, Cherry S, Sadovsky Y, Coyne CB. 2016. Type III interferons produced by human placental trophoblasts confer protection against Zika virus infection. Cell Host Microbe 19:705–712. doi: 10.1016/j.chom.2016.03.008 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B45] 45. Coetzer JA, Barnard BJ. 1977. Hydrops amnii in sheep associated with hydranencephaly and arthrogryposis with wesselsbron disease and rift valley fever viruses as aetiological agents. Onderstepoort J Vet Res 44:119–126. [PubMed] [Google Scholar]

[B46] 46. Bird BH, Albariño CG, Nichol ST. 2007. Rift Valley fever virus lacking NSM proteins retains high virulence in vivo and may provide a model of human delayed onset neurologic disease. Virology 362:10–15. doi: 10.1016/j.virol.2007.01.046 [DOI] [PubMed] [Google Scholar]

[B47] 47. Billecocq A, Spiegel M, Vialat P, Kohl A, Weber F, Bouloy M, Haller O. 2004. Nss protein of Rift Valley fever virus blocks interferon production by inhibiting host gene transcription. J Virol 78:9798–9806. doi: 10.1128/JVI.78.18.9798-9806.2004 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B48] 48. Koenig ZT. 2018. Type III interferon control of Rift Valley fever virus at epithelial cell barrier. Master of Science. University of Pittsburgh, SUNY Geneseo. [Google Scholar]

PERMALINK

Vaccine strains of Rift Valley fever virus exhibit attenuation at the maternal–fetal placental interface

Cynthia M McMillen

Christina Megli

Rebecca Radisic

Lauren B Skvarca

Ryan M Hoehl

Devin A Boyles

Jackson J McGaughey

Brian H Bird

Anita K McElroy

Amy L Hartman

Roles

ABSTRACT

IMPORTANCE

INTRODUCTION

RESULTS

Fig 1.

Sheep placenta explants are permissive to infection with RVFV vaccine strains

TABLE 1.

Fig 2.

Fig 3.

All vaccine strains infect human villous explants, but tissue distribution differs in viruses lacking the viral NSs protein

TABLE 2.

Fig 4.

Fig 5.

RVFV NSs antagonizes production of both type I and III interferons in human villous cultures

Fig 6.

Sheep placenta explant cultures produce more virus than human placenta explant cultures

Fig 7.

DISCUSSION

MATERIALS AND METHODS

Viruses

Biosafety

Placenta procurement

Explant studies

RNA isolation and q-RT-PCR

Enzyme-linked immunosorbent assay

Immunohistochemistry and microscopy

Statistics

ACKNOWLEDGMENTS

Contributor Information

ETHICS APPROVAL

DATA AVAILABILITY

SUPPLEMENTAL MATERIAL

REFERENCES

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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