Rift Valley Fever Virus

Molecular Pathogenesis: Viral Entry, NSs-Mediated Immune Evasion, and Host Factors

The molecular pathogenesis of Rift Valley fever virus (RVFV) is a sophisticated interplay between viral determinants of entry and replication, a multifaceted arsenal of host immune evasion strategies, and a diverse array of host factors that either restrict or facilitate infection. Understanding this intricate molecular dialogue is paramount for rational development of vaccines and therapeutics against this WHO-priority and CDC/NIAID Category A pathogen, which continues to expand its geographic range beyond Africa and the Arabian Peninsula [1, 4, 5].

Viral Entry Mechanisms: A Multistep Orchestration

The initial phase of RVFV infection is governed by the viral envelope glycoproteins, Gn and Gc, which are arranged as heterodimers forming higher-order oligomers on the virion surface [6, 12, 14]. The Gn glycoprotein serves as the primary attachment factor, engaging host cell receptors to initiate clathrin-mediated endocytosis. A landmark genome-wide CRISPR screen identified low-density lipoprotein receptor-related protein 1 (LRP1) as a critical host entry factor for RVFV [2]. This study demonstrated that RVFV Gn directly binds to specific clusters on LRP1 in a glycosylation-independent manner, and that exogenous addition of the murine receptor-associated protein domain 3 (mRAPD3) or anti-Lrp1 antibodies potently neutralizes infection across diverse cell lines. The therapeutic potential of targeting this interaction was confirmed in vivo, where mRAPD3 treatment protected mice from lethal RVFV challenge [2]. Heat shock protein Grp94 was also identified as a critical co-factor in this screen, likely assisting in proper folding or trafficking of LRP1 or the viral glycoproteins [2].

Beyond LRP1, dendritic cell-specific ICAM-3 grabbing non-integrin (DC-SIGN) functions as an alternative attachment factor, particularly relevant for infection of dendritic cells and macrophages. This interaction is mediated by N-glycans on the Gn and Gc glycoproteins, which were shown to redundantly support viral infection via DC-SIGN [24]. Specifically, N-glycans at positions N438 of Gn and N1077 of Gc facilitate this interaction, although other Gc N-glycans (N794, N1035) also contribute [24]. This redundancy suggests that RVFV has evolved to utilize multiple entry routes, enhancing its tropism for critical immune cell populations.

Following attachment, the low pH environment of the endosome triggers dramatic conformational rearrangements in the Gn-Gc complex. This process, which can be blocked by potent neutralizing antibodies, exposes the class II fusion loop in Gc, driving membrane fusion and release of the viral ribonucleocapsid into the cytoplasm [6, 9, 12]. Structural studies have revealed that the membrane-distal domain of Gn, particularly domain III, constitutes a major site of vulnerability targeted by neutralizing antibodies that prevent these fusogenic rearrangements [9, 12]. The human antibody response following natural infection or vaccination is dominated by Gn-specific antibodies, which correlate strongly with neutralizing titers and protective immunity [6, 11]. This underscores Gn as the primary target for both humoral immunity and rational vaccine design [17, 21].

NSs-Mediated Immune Evasion: A Masterclass in Host Subversion

The non-structural protein NSs, encoded by the S-segment, is the major virulence factor of RVFV, orchestrating a comprehensive shutdown of the host antiviral response [3, 15]. This approximately 29 kDa protein operates through at least three distinct, yet interconnected, mechanisms that collectively render infected cells profoundly susceptible to viral replication.

Interferon Antagonism via Transcriptional Suppression: NSs potently inhibits host cell transcription, including the expression of type I interferon (IFN) genes. It achieves this by interacting with the p62 subunit of the TFIIH transcription factor complex, targeting it for proteasomal degradation [15]. By eliminating p62, NSs disrupts RNA polymerase II-mediated transcription, effectively silencing the expression of hundreds of host genes, including IFN-β. Furthermore, NSs forms a complex with the Sin3A-associated protein 30 (SAP30) and the transcription factor YY1, bridging this complex to chromatin DNA [15]. This NSs-SAP30-YY1 interaction interferes with chromatin segregation and cohesion and is essential for the specific suppression of IFN-β promoter activation, providing an additional layer of control over the innate immune response.

Degradation of Protein Kinase R (PKR): The antiviral kinase PKR is a central effector of the IFN response, halting viral protein synthesis by phosphorylating eIF2α upon detection of double-stranded RNA. RVFV NSs eliminates this potent restriction factor by co-opting the host ubiquitin-proteasome system. NSs assembles a specialized SCF (SKP1-CUL1-F-box) E3 ubiquitin ligase complex by recruiting the F-box proteins FBXW11 and, to a lesser extent, β-TrCP1 [18, 20]. Through a critical six-amino-acid degron sequence (DDGFVE) within NSs, the protein bridges the SCFFBXW11 complex with PKR, facilitating its polyubiquitination and rapid proteasomal degradation [18]. This degradation is essential for RVFV replication; disruption of this interaction, through siRNA knockdown of FBXW11, by using a small molecule inhibitor of SCF ligase activity (MLN4924), or by introducing degron mutations, stabilizes PKR, leading to its activation and potent suppression of viral translation [18]. The redundancy of utilizing both FBXW11 and β-TrCP1 underscores the critical importance of PKR elimination for RVFV fitness [20].

Formation of Nuclear and Cytosolic Amyloid-Like Fibrils: A truly remarkable and recently elucidated aspect of NSs function is its ability to self-assemble into amyloid-like fibrils. These structures, visualized in the nuclei and cytoplasm of infected cells and in the brains of infected mice, exhibit classic amyloid properties: a 12 nm width, strong detergent resistance, and binding to Thioflavin-S [8]. The formation of these NSs filaments is dependent on disulfide bonds and occurs spontaneously within hours of infection. Crucially, the presence of these amyloid-like aggregates was directly associated with increased mortality in a mouse model of RVFV encephalitis [8]. This represents a paradigm-shifting discovery: a viral protein can encode a structural motif that forms pathogenic amyloid aggregates, potentially contributing to neuropathology and tissue damage through mechanisms distinct from its established immune evasion functions. This aggregation may also sequester host proteins or disrupt nuclear architecture, further compounding the cellular damage [15].

The cumulative effect of these NSs functions is a profound inhibition of both the induction (via transcriptional suppression) and the effector phases (via PKR degradation) of the innate immune response. Mutations or deletions within the NSs gene, as seen in vaccine candidates like Clone 13 or ΔNSs constructs, render the virus highly attenuated, allowing for the induction of a robust protective immune response [3, 16, 25].

Host Factors and Restriction: The Intracellular Battlefield

The outcome of RVFV infection is also profoundly shaped by host factors that either promote or restrict viral replication. These host-virus interactions represent critical nodes for pathogenesis and potential therapeutic intervention.

Restriction by Innate Immune Sensors and Effectors: Beyond the pathways targeted by NSs, host cells employ multiple layers of antiviral defense. The DEAD-box helicase DDX17 has been identified as a restriction factor that recognizes and unwinds the non-coding regions of the RVFV S-segment, namely the intergenic region (IGR) and the 5' non-coding region (NCR) [10]. Biochemical studies confirmed that the DDX17 helicase domain binds to these structured RNAs with micromolar affinity and possesses ATP-dependent unwinding activity, suggesting a direct antiviral mechanism [10]. In the central nervous system (CNS), the mitochondrial antiviral-signaling protein (MAVS) is critical for orchestrating a protective immune response in microglia. MAVS-deficient mice exhibited enhanced susceptibility to RVFV encephalitis, characterized by increased brain viral burden and mortality, due to defective type I interferon and interferon-stimulated gene (ISG) expression in microglia and dysregulated lymphocyte infiltration [7]. This highlights the importance of the RIG-I/MAVS pathway as a frontline defense in the brain, a key target tissue for severe RVF.

Proviral Host Factors and Pathways: Conversely, RVFV commandeers a variety of cellular pathways to support its replication. A genome-wide RNA interference screen identified the canonical Wnt/β-catenin signaling pathway as a key proviral pathway [23]. RVFV infection activates this pathway, and its enhancement promotes infection, while pharmacological inhibition or genetic knockdown of β-catenin reduces viral replication. This dependency on Wnt signaling is shared by other bunyaviruses, suggesting a conserved mechanism to regulate cell cycle conditions conducive to viral replication [23]. Another critical proviral factor is serine/threonine protein phosphatase 1 (PP1). Pharmacological inhibition of PP1 or its knockdown significantly reduces RVFV replication early in the life cycle, impacting viral RNA production, likely through interaction with the viral L-polymerase and nucleoprotein [26]. Furthermore, RVFV triggers the release of exosomes that contain viral RNA (L, M, and S segments) and viral proteins, including N and a modified form of NSs [19]. These virus-modified exosomes can induce apoptosis in recipient immune cells, suggesting a mechanism for immune subversion and systemic pathogenesis that extends beyond direct cell-to-cell spread [19].

Antiviral Defenses in the Arthropod Vector: In mosquitoes, the primary antiviral defense is the RNA interference (RNAi) pathway. RVFV infection in vector species such as Aedes and Culex induces the production of virus-derived small interfering RNAs (siRNAs) and Piwi-interacting RNAs (piRNAs), which guide the RNA-induced silencing complex to degrade viral RNA [13]. This pathway is a significant barrier to viral replication in the insect host, and its suppression by the virus is critical for vector competence. Notably, RVFV does not appear to encode a dedicated suppressor of RNAi, unlike many plant and insect-specific viruses [13]. This suggests that the virus relies on high replication rates and exploitation of vector-specific host factors to overcome this innate immune barrier, a dynamic that is crucial for understanding viral maintenance during inter-epidemic periods [22] and spillover events.

Epidemiology and Vector-Borne Transmission Dynamics

Rift Valley fever virus (RVFV) is a mosquito-borne phlebovirus that represents one of the most significant emerging arboviral threats to both human and animal health across Africa and the Arabian Peninsula [1, 5]. The virus is classified as a Category A pathogen by the U.S. Centers for Disease Control and Prevention (CDC) and the National Institute of Allergy and Infectious Diseases (NIAID), and is included among the eight priority pathogens on the World Health Organization (WHO) Blueprint list, underscoring its pandemic potential and the urgent need for comprehensive surveillance and control strategies [1, 19]. The epidemiology of RVFV is characterized by a complex interplay between mosquito vectors, vertebrate reservoir and amplifying hosts, environmental drivers, and anthropogenic factors that together determine the spatial and temporal patterns of viral transmission and disease emergence.

Geographic Distribution and Historical Emergence

Since its initial identification in 1931 during an outbreak among sheep in the Rift Valley region of Kenya, RVFV has demonstrated a remarkable capacity for geographic expansion and re-emergence [5, 11]. Phylogenetic analyses suggest that the virus first emerged in the mid-19th century, with subsequent diversification and spread across the African continent [5]. The virus is now endemic throughout much of sub-Saharan Africa, with documented circulation in at least 39 countries between 1999 and 2021 alone [4]. Major epizootics and epidemics have occurred with increasing frequency and severity, including the devastating 1977 outbreak in Egypt that resulted in an estimated 200,000 human infections and 600 deaths, the 1987 outbreak in West Africa, and the 2000 emergence in the Arabian Peninsula, marking the first documented occurrence of RVFV outside of Africa [5, 40].

The phylogeographic history of RVFV reveals a dynamic pattern of viral movement and diversification. Phylogeographic analyses indicate that East Africa, particularly Kenya, serves as a major source population from which viral lineages have repeatedly disseminated to other regions [40]. Madagascar experienced at least three independent introductions: the first from Zimbabwe, followed by two subsequent introductions from Kenya [40]. The Egyptian outbreaks of 1977 and subsequent years likely originated from a long-distance introduction from Zimbabwe, while the Arabian Peninsula outbreak resulted from a single introduction from Kenya [40]. West African viral populations appear to have arisen from a single introduction from the Central African Republic, with subsequent bidirectional movement observed between Kenya and Sudan, as well as between the Central African Republic and Zimbabwe [40]. This extensive mobility underscores the invasive potential of RVFV and the constant threat of introduction into previously unaffected regions.

Vector Species and Vector Competence

RVFV is transmitted by a diverse array of mosquito species, with over 50 species demonstrated to be competent vectors under laboratory conditions [5, 30]. The primary vectors belong to the genera Aedes and Culex, although species within Anopheles, Mansonia, and Coquillettidia have also been implicated in transmission [1, 37, 39]. The vector competence of different mosquito species varies considerably depending on viral strain, mosquito genotype, environmental conditions, and experimental methodology [31]. A comprehensive meta-analysis of Mediterranean mosquito species identified Aedes caspius as potentially the most competent vector among five major species evaluated, including Ae. detritus, Ae. vexans, Culex pipiens, and Cx. theileri [31]. However, the vector competence of Cx. pipiens and Ae. albopictus has been confirmed for European populations, raising concerns about the potential for autochthonous transmission should the virus be introduced into Europe [35].

The extrinsic incubation period (EIP), the time between ingestion of an infectious blood meal and the ability to transmit virus via saliva, is a critical determinant of vectorial capacity. For RVFV, the EIP is temperature-dependent, with higher temperatures generally accelerating viral dissemination and transmission [45]. Studies using European Cx. pipiens and Ae. detritus maintained at 20°C or 25°C demonstrated that transmission efficiency, while present, was relatively low even at elevated temperatures, with average transmission rates of 5-7% [45]. This finding suggests that while temperate mosquitoes are competent vectors, environmental conditions may limit the efficiency of transmission in cooler climates.

Vertical Transmission and Inter-Epidemic Maintenance

One of the most critical and long-debated aspects of RVFV epidemiology is the mechanism by which the virus persists during inter-epidemic periods (IEPs), which can last 4-10 years or longer [22, 44]. Vertical transmission, the passage of virus from infected female mosquitoes to their offspring, has been proposed as a key mechanism for viral maintenance during dry periods when adult mosquito populations are low. For decades, this phenomenon remained unconfirmed in the laboratory despite strong epidemiological evidence, including the ability to predict RVFV epizootics based on rainfall patterns derived from satellite data [27].

In 2021, Bergren and colleagues provided the first laboratory confirmation of vertical transmission of RVFV by Culex tarsalis mosquitoes following oral exposure [27]. Progeny from three successive gonotrophic cycles were reared to adulthood, with infectious RVFV confirmed in each developmental stage. Virus was detected in ovarian tissues of parental mosquitoes 7 days after imbibing an infectious blood meal, and infection rates among progeny ranged from 2.0-10.0% [27]. Importantly, viral titers among progeny were low, suggesting the presence of host mechanisms that suppress viral replication, potentially limiting the efficiency of this transmission route but still providing a mechanism for viral persistence [27].

The role of vertical transmission in RVFV ecology is further supported by field observations. During the 2016 outbreak in Uganda, the first laboratory-confirmed outbreak in 48 years, entomological surveys detected RVFV RNA in pools of Aedes and Coquillettidia species, with 1% of pools testing positive [37]. The detection of virus in these vectors during an active outbreak, combined with the known ability of Aedes mosquitoes to transmit virus transovarially, supports the hypothesis that infected mosquito eggs serve as a reservoir for RVFV during inter-epidemic periods [22, 37].

However, recent evidence suggests that vertical transmission alone may not be sufficient to explain viral persistence. Njenga and Bett (2018) reviewed accumulating evidence indicating that low-level cycling of RVFV among wildlife, livestock, and humans occurs during IEPs, maintained by mosquito vectors in ecosystems characterized by low annual rainfall and specific soil types [22]. This hypothesis is supported by numerous serological studies documenting ongoing viral circulation during IEPs. In Mozambique, seroprevalence rates of 44.2% in sheep and 25.1% in goats were observed in the absence of reported outbreaks, indicating subclinical inter-epidemic transmission [49]. Similarly, a longitudinal study in Kenya documented 15 seroconversions in small ruminants within irrigated and riverine villages during an IEP, while no seroconversions were detected in pastoral ecosystems [44]. These findings highlight the importance of environmental modification, particularly irrigation, in creating conditions that support vector breeding and viral transmission during periods that would otherwise be unfavorable for transmission [44].

Transmission Cycles: Sylvatic, Epizootic, and Epidemic

RVFV transmission occurs through three interconnected cycles: a sylvatic (enzootic) cycle involving wildlife and primarily Aedes mosquitoes, an epizootic cycle involving livestock and various mosquito species, and an epidemic cycle involving humans [5, 22]. The sylvatic cycle is thought to maintain the virus in nature, with vertical transmission in Aedes mosquitoes serving as the primary mechanism for viral persistence during inter-epidemic periods [22, 27]. When environmental conditions become favorable, typically following periods of heavy rainfall and flooding, Aedes eggs hatch, and infected mosquitoes emerge to initiate transmission to susceptible vertebrate hosts [5, 39].

The transition from sylvatic to epizootic transmission is driven by the amplification of virus in livestock populations. Sheep, cattle, and goats develop high-titer viremia sufficient to infect feeding mosquitoes, thereby amplifying transmission [5, 30, 48]. The experimental reproduction of the mosquito-lamb-mosquito transmission cycle by Schreur and colleagues demonstrated that feeding mosquitoes on viremic lambs resulted in strikingly higher infection rates compared to artificial membrane feeding, highlighting the importance of using natural host-vector systems for understanding transmission dynamics [30]. Once established in livestock populations, secondary vectors, particularly Culex species, can sustain and expand the outbreak, as these mosquitoes are often more abundant and have different feeding preferences than the primary Aedes vectors [5, 39].

Human infection occurs primarily through two routes: mosquito bites and direct contact with infected animal tissues or bodily fluids [1, 36]. The relative contribution of each route varies by epidemiological context. During epizootics, the majority of human cases are thought to result from mosquito transmission, particularly in areas with high vector densities [5]. However, occupational exposure, including slaughtering, butchering, assisting with animal births, and handling aborted materials, represents a significant risk factor for infection [4, 36]. A systematic review and meta-analysis by Gerken and colleagues found that animal contact, butchering, milking, and handling aborted material were significantly associated with greater odds of RVFV exposure [4].

Risk Factors for Human Infection

The epidemiology of human RVFV infection is characterized by distinct demographic and behavioral risk factors. Men have consistently been found to have greater odds of RVFV infection than women, likely reflecting occupational exposure to livestock and animal products [4]. However, women may be at increased risk through domestic activities such as milking and processing animal products [36]. A study of raw milk exposures in Kenya found that 77.2% of seropositive individuals engaged in milking livestock compared to 32.0% of seronegative individuals, and 86.5% of seropositive individuals consumed raw milk compared to 33.4% of seronegative individuals [36]. The odds of RVFV exposure were significantly higher for individuals who both milked and consumed raw milk compared to those whose only contact was through milking, suggesting that ingestion of contaminated milk may represent a previously underappreciated transmission route [36].

The association between RVFV infection and adverse pregnancy outcomes represents a particularly concerning aspect of the virus's epidemiology. Baudin and colleagues conducted a cross-sectional study of febrile pregnant women in Sudan and found that 54% of women with acute RVFV infection experienced miscarriage, compared to 12% of RVFV-negative women [38]. Multivariate logistic regression identified acute RVFV infection as an independent predictor of miscarriage, with an odds ratio of 7.4 [38]. This finding is consistent with experimental studies demonstrating that RVFV targets the maternal-fetal interface in both ovine and human placentas [29]. In pregnant ewes, the virus replicates efficiently in maternal placental epithelial cells before infecting fetal trophoblasts, ultimately resulting in placental and fetal demise followed by abortion [29]. The virus can bypass the maternal epithelial cell layer by directly targeting fetal trophoblasts in the hemophagous zone, a region of the ovine placenta where maternal blood is in direct contact with fetal cells [29]. Experiments with human placental explants confirmed that RVFV replicates efficiently in both cytotrophoblasts and syncytiotrophoblasts, underscoring the risk of RVFV infection for human pregnancy [29].

Environmental Drivers and Climate Change

The epidemiology of RVFV is profoundly influenced by environmental factors, particularly rainfall, temperature, and humidity, which affect vector population dynamics and viral transmission efficiency [5, 39, 43]. Outbreaks are typically associated with periods of heavy rainfall and flooding, which create extensive breeding sites for mosquitoes and trigger the hatching of infected Aedes eggs [5, 39]. Ecological niche modeling has identified precipitation in the driest quarter, precipitation seasonality, and isothermality as the most important predictors of RVFV vector distribution [50]. Soil type also plays a critical role, with certain soil types being more favorable for mosquito egg survival and larval development [22, 50].

The impact of climate change on RVFV epidemiology is a growing concern. As global temperatures rise and precipitation patterns shift, the geographic range of competent vectors is expected to expand, potentially introducing the virus into previously unaffected regions [1, 5, 41]. Ecological niche modeling for Baringo County, Kenya, predicted that under future climatic conditions (2050), the spatial distribution of Culex quinquefasciatus and Mansonia africana would increase, expanding the areas suitable for RVFV transmission [50]. Similarly, the presence of competent Cx. pipiens and Ae. albopictus in Europe, combined with warming temperatures, raises the possibility of autochthonous RVFV transmission should the virus be introduced [35, 41].

The role of irrigation in modifying local environmental conditions and facilitating RVFV transmission during inter-epidemic periods deserves particular attention. In Kenya, seroconversions were detected only in irrigated and riverine villages, not in pastoral ecosystems, suggesting that irrigation provides necessary environmental conditions that enable vectors access to more breeding grounds, resting places, and shade, favoring their breeding and survival [44]. This finding has important implications for land-use planning and vector control strategies in endemic areas.

Livestock and Wildlife as Amplifying Hosts

Livestock, particularly sheep, cattle, and goats, serve as the primary amplifying hosts for RVFV, developing high-titer viremia sufficient to infect feeding mosquitoes [5, 30, 48]. The susceptibility and clinical outcome of infection vary by species and age. Sheep are considered the most susceptible, with mortality rates approaching 100% in neonates and abortion rates of nearly 100% in pregnant ewes [5, 14]. Cattle are also highly susceptible, although clinical disease may be less severe than in sheep [48]. Experimental infection of calves with two genetically distinct RVFV strains (Kenya 2006 and Saudi Arabia 2001) demonstrated that the Kenyan strain caused more severe disease, with higher viremia titers, more pronounced liver pathology, and elevated liver enzyme levels [48]. This finding highlights the importance of viral genetic diversity in determining pathogenicity and transmission potential.

Camels have emerged as an increasingly important livestock species in the epidemiology of RVFV, particularly in arid and semi-arid regions where they are becoming the livestock of choice for pastoralists adapting to climate change [28]. A serosurvey in northern Kenya found that 14.2% of dromedary camels were seropositive for RVFV, with 27.5% seropositive for at least one of three zoonotic pathogens (RVFV, Brucella spp., or Coxiella burnetii) [28]. The high seropositivity rates indicate the endemicity of RVFV among camel populations and suggest that camels may play a significant role in viral transmission dynamics, particularly in areas where they are the primary livestock species.

The role of wildlife in RVFV maintenance and transmission is less well understood but increasingly recognized as important. Serological evidence of RVFV infection has been documented in various wildlife species, including African buffalo, elephants, rhinoceroses, and various antelope species [32, 42]. Notably, neutralizing antibodies against RVFV have been detected in Ugandan bats, with 9.6% of little epauletted fruit bats (Epomophorus labiatus) and 5.6% of Egyptian rousette bats (Rousettus aegyptiacus) testing positive [42]. The potential role of bats as reservoir hosts warrants further investigation, particularly given their ability to harbor other zoonotic viruses and their extensive geographic range.

Spatial and Temporal Patterns of Transmission

The transmission of RVFV exhibits distinct spatial and temporal patterns that reflect the complex interactions between vectors, hosts, and environmental factors. Outbreaks occur at irregular intervals, typically every 4-15 years, and are often associated with El Niño-Southern Oscillation (ENSO) events that bring heavy rainfall to East Africa [5, 47]. However, the frequency of outbreaks has increased in recent decades, with human epidemics occurring in 1998, 2006-2007, and 2008 in East Africa alone [47]. This increasing frequency may reflect both improved surveillance and true changes in transmission dynamics driven by climate change, land-use change, and viral evolution [47].

Spatially, RVFV transmission is heterogeneous, with certain areas experiencing recurrent outbreaks while others remain relatively unaffected. In Africa, Mauritania, Madagascar, Kenya, South Africa, and Sudan have reported the most human outbreak years between 1999 and 2021 [4]. Within these countries, transmission is often concentrated in specific ecosystems, such as the Senegal River Delta and Valley in Senegal, where ecological conditions favor high vector densities [39]. A study in Senegal found that RVFV vectors represented 89.02% of all mosquitoes captured, with Ae. vexans arabiensis (31.29%), Cx. tritaeniorhynchus (33.09%), and Ma. uniformis (24.04%) being the most abundant species [39]. The abundance of these vectors was significantly influenced by temperature, relative humidity, rainfall, and the type of biotope (temporary ponds, rivers, or lakes) [39].

Mathematical modeling has provided important insights into the spatial dynamics of RVFV transmission. A metapopulation model developed for the Comoros archipelago demonstrated that the archipelago network was able to sustain viral transmission in the absence of explicit disease introduction events after early 2007, and that repeated outbreaks during 2004-2020 may have gone under-detected by local surveillance [34]. The model also showed that coordinated within-island control measures are more effective than between-island animal movement restrictions, highlighting the importance of local vector control and livestock management [34].

Viral Genetic Diversity and Evolution

The genetic diversity of RVFV has important implications for its epidemiology and transmission dynamics. The virus has a tripartite, single-stranded RNA genome consisting of small (S), medium (M), and large (L) segments [5, 14, 33]. The segmented nature of the genome allows for reassortment, the exchange of genome segments between different viral strains, which can generate novel variants with altered pathogenicity and transmission characteristics [5, 47]. Phylogenetic analyses have identified multiple lineages of RVFV, with the Kenya-2 clade being responsible for the 2006-2007 East African outbreak and the 2016 Ugandan outbreak [37, 47].

The accumulation of genetic mutations over time has been associated with increasing severity of human disease in East Africa [47]. Comparisons between isolates from different outbreaks have revealed an accumulation of genetic mutations and genomic reassortments that have diversified RVFV genomes over several decades [47]. The Northeastern African lineage, which was responsible for the 2015 outbreak in Mauritania, represents a distinct genetic group that originated from Northeastern Africa and has been associated with severe human disease [46]. During this outbreak, 57 confirmed cases and 12 deaths were reported, with hemorrhagic manifestations observed in

Clinical Manifestations and Diagnostic Challenges

Rift Valley fever virus (RVFV) presents a formidable diagnostic conundrum precisely because its clinical manifestations span an extraordinarily broad spectrum, ranging from a completely asymptomatic seroconversion to a fulminant, fatal hemorrhagic syndrome. This pleomorphism is not merely a taxonomic curiosity; it is the central obstacle to timely case identification, outbreak containment, and effective patient management. The virus, a mosquito-borne phlebovirus in the order Bunyavirales, is recognized by the World Health Organization (WHO) as a priority pathogen on its Bluepoint list, a designation that underscores its pandemic potential and the critical gaps in our diagnostic armamentarium [1, 5]. The clinical challenge is further compounded by the fact that the majority of human infections, estimated at upwards of 80%, are either asymptomatic or present as a non-specific, acute febrile illness that is clinically indistinguishable from malaria, dengue, chikungunya, or typhoid fever [1, 3, 49, 59]. This diagnostic ambiguity, particularly in the early stages of infection, provides a critical window for unchecked viral dissemination and delays the implementation of public health interventions.

The Spectrum of Human Disease: From Undifferentiated Fever to Multi-Organ Failure

In the symptomatic individual, the incubation period typically ranges from two to six days following exposure via mosquito bite or contact with infected animal tissues or bodily fluids [1, 3, 36]. The initial prodrome is characterized by the abrupt onset of fever, headache, myalgia, arthralgia, and lassitude, a constellation of symptoms so generic that it is often dismissed in endemic settings [1, 49]. However, a minority of patients, approximately 1-2% of all infections, will progress to one of three severe, life-threatening manifestations: hemorrhagic fever, encephalitis, or ocular disease [3, 51, 61]. The hemorrhagic form, which carries the highest case-fatality rate, typically emerges 2-4 days after the onset of illness and is marked by jaundice, petechiae, ecchymoses, epistaxis, hematemesis, melena, and gingival bleeding [3, 46, 61]. Laboratory findings in these patients consistently reveal severe thrombocytopenia, leukopenia, anemia, and markedly elevated serum transaminases (AST and ALT), reflecting massive hepatocellular necrosis [3, 48, 61]. Indeed, hepatic involvement is a hallmark of severe RVFV infection, and the degree of liver enzyme elevation often correlates with disease severity and outcome [3, 48]. The pathogenesis of this hepatic phase is driven by the virus's robust replication in hepatocytes, leading to disseminated intravascular coagulation (DIC) and multi-organ failure [3, 5]. The 2015 outbreak in southern Mauritania, documented by Boushab et al., provided a stark illustration of this syndrome: among 31 confirmed cases, 81% presented with hemorrhagic manifestations, and the overall case-fatality rate reached 42%, with all fatalities exhibiting hemorrhagic disease [46, 61].

Neurological and Ocular Involvement: The Delayed Encephalitic Phase

A unique and particularly challenging aspect of RVFV pathogenesis is the late-onset encephalitic syndrome, which can appear days to weeks after the initial febrile illness has seemingly resolved [51-53]. This neurological manifestation, which occurs in less than 1% of infected individuals, is characterized by severe headache, confusion, hallucinations, vertigo, neck stiffness, ataxia, seizures, and focal motor deficits, including hemiparesis and cranial nerve palsies [3, 51, 53]. The virus exhibits a pronounced neurotropism, and the mechanisms by which it invades the central nervous system (CNS) remain an area of active investigation. Data from animal models, including the newly characterized CC057/Unc strain of Collaborative Cross mice and the ferret model, indicate that RVFV can enter the CNS via olfactory or hematogenous routes, subsequently replicating to high titers in the brain parenchyma [7, 51-53]. Importantly, this encephalitic phase is frequently associated with viral clearance from the periphery, meaning that diagnostic tests based on blood samples, such as RT-PCR, may be falsely negative by the time neurological symptoms appear [52, 53, 57]. The ferret model demonstrated that animals with clinical CNS disease (seizures, ataxia, hind limb weakness) had high viral RNA loads in brain tissue despite transient or absent viremia [53]. Furthermore, the NSs protein, the major virulence factor of RVFV, has been shown to form amyloid-like fibrils in the brains of infected mice, a finding that may contribute to the neuropathology and long-term sequelae of this form of the disease [8]. Ocular complications, including retinitis, macular degeneration, and temporary or permanent vision loss, can also occur and may be the only manifestation of infection in some patients [3, 51]. The retina is a direct target of RVFV, and ocular disease can present concurrently with or independently of other severe outcomes.

The Unique Challenge of RVFV in Pregnancy: Abortion and Vertical Transmission

Perhaps the most epidemiologically and clinically significant manifestation of RVFV, one that is well-recognized in livestock but increasingly documented in humans, is its abortigenic potential. In ruminants, RVFV infection of pregnant animals is infamous for causing "abortion storms," where up to 100% of pregnant ewes, cows, or goats may abort, often without the dam showing any other clinical signs of illness [3, 29, 30, 58]. The pathological basis for this catastrophic outcome has been elucidated in recent years. Oymans et al. demonstrated that RVFV targets the maternal-fetal interface in both ovine and human placentas, replicating efficiently in maternal placental epithelial cells before infecting fetal trophoblasts [29]. The virus can bypass the maternal epithelial barrier entirely by directly targeting fetal cells in the hemophagous zone of the ovine placenta, leading to widespread necrosis, hemorrhage, and fetal demise [29]. McMillen et al. replicated these findings in a pregnant rat model, showing that RVFV infection results in intrauterine fetal death and severe congenital abnormalities even in dams that exhibit no signs of illness [54]. Critically, Baudin et al. provided the first direct evidence linking RVFV infection to miscarriage in pregnant women in Sudan, finding that 54% of febrile pregnant women with acute RVFV infection experienced miscarriage, compared to just 12% of RVFV-negative febrile women, yielding an adjusted odds ratio of 7.4 [38]. This finding has profound implications for public health messaging and clinical management during outbreaks, yet the diagnosis of RVFV-associated miscarriage remains challenging because the mother may be asymptomatic or present with only mild fever.

Diagnostic Challenges: The Narrow Window of Detectability

The clinical diagnosis of RVFV infection is notoriously difficult for several interconnected reasons. First, as detailed above, the early symptoms are indistinguishable from a host of other endemic febrile illnesses, meaning that clinical acumen alone is insufficient for accurate diagnosis [1, 59]. Second, the virus is detectable in the blood only for a very brief period, typically 3-5 days following the onset of fever, after which it is cleared from the periphery by the developing immune response [1, 52, 53]. This transient viremia means that molecular diagnostic methods, such as real-time reverse transcription polymerase chain reaction (RT-PCR), have a very narrow window of utility. If a patient presents later in the course of illness, for example, with encephalitis or hemorrhagic fever, their blood may be RT-PCR-negative, even though the virus is actively replicating in the brain or other tissues [1, 52]. This diagnostic "blind spot" is a major impediment to both clinical care and epidemiological surveillance.

Consequently, there is a broad consensus that reliance on a single diagnostic modality is inadequate. The optimal approach for case determination involves the strategic combination of molecular and serological testing [1, 41]. During the acute phase (days 1-5), RT-PCR or virus isolation from serum is the method of choice. After the first week, serological methods, specifically IgM-capture enzyme-linked immunosorbent assay (ELISA) for recent infection and IgG ELISA for past exposure, become essential [1, 41, 56]. However, serological testing is not without its own challenges. The potential for cross-reactivity with other phleboviruses (e.g., Toscana virus, sandfly fever viruses) can confound results, and the kinetics of the antibody response can vary significantly between individuals [1, 25]. Furthermore, immune-compromised patients or those with rapidly fatal disease may not mount a detectable antibody response before succumbing to infection [1].

The lack of a validated, point-of-care diagnostic test is a critical gap in our ability to manage RVFV outbreaks [1]. Current diagnostic workflows require sample transport to centralized, high-containment (BSL-3/BSL-4) laboratories, which is logistically impractical in many endemic regions of sub-Saharan Africa, where health systems are often fragile and laboratory infrastructure is limited [1, 14]. This delay between sample collection and result generation can span days, a period during which the index patient may transmit the virus to caregivers or family members, and the public health response is delayed.

Subclinical Infection and Inter-Epidemic Persistence

A further diagnostic challenge, and one that profoundly influences our understanding of RVFV epidemiology, is the high prevalence of subclinical or undifferentiated infections, particularly during inter-epidemic periods (IEPs). Numerous serosurveys have demonstrated that RVFV circulates continuously at low levels in both human and livestock populations, even in the absence of recognized outbreaks [22, 32, 44, 55, 56, 60]. For example, in KwaZulu-Natal, South Africa, Bergh et al. observed an annual seroconversion rate of 0.59 per animal-year in cattle and 0.41 per animal-year in goats, with seroprevalence exceeding 30% in some localities, despite no reported outbreaks [55]. Similarly, in Mozambique, Mbotha et al. documented inter-epidemic seroconversions in sheep and goats associated with irrigated agricultural systems [44]. These findings highlight that the virus is not simply "hibernating" in mosquito eggs during long dry spells, as was once hypothesized, but is actively maintained through low-level cycling between vectors and vertebrates [22, 44]. This cryptic circulation poses a major diagnostic surveillance challenge: standard passive surveillance systems, which rely on the reporting of clinical signs (abortion storms, hemorrhagic disease), are insensitive to this low-level activity. Active surveillance, combining sentinel animal serology and molecular screening of mosquito populations, is required to detect the virus before it can amplify into an explosive epizootic [4, 22, 34]. The WHO and the Food and Agriculture Organization (FAO) have consistently called for integrated human-animal-vector surveillance to close this diagnostic gap, but implementation remains fragmentary and underfunded.

In summary, the clinical manifestations of RVFV infection are profoundly diverse, ranging from inapparent infection and non-specific febrile illness to devastating hepatitis, hemorrhagic fever, encephalitis, and reproductive failure. This spectrum, combined with the transient nature of viremia and the absence of point-of-care diagnostics, creates a uniquely challenging diagnostic landscape. The clinician must maintain a high index of suspicion in febrile patients from endemic areas, particularly those with known livestock contact, and must employ a combined molecular and serological testing strategy that is tailored to the stage of illness. The failure to diagnose RVFV in a timely manner has direct consequences: it permits continued transmission, hinders outbreak control, and deprives patients of potentially life-saving supportive care. The development of rapid, multiplex diagnostic platforms that can simultaneously differentiate RVFV from other hemorrhagic fever viruses and arboviruses remains a priority of the highest order for global health security.

Advanced Diagnostics: Molecular and Serological Approaches

The accurate and timely diagnosis of Rift Valley fever virus (RVFV) infection is a cornerstone of effective outbreak response, surveillance, and patient management. The diagnostic landscape for RVFV is complex, shaped by the virus's transient viremia, the non-specific nature of early clinical presentations, and the critical need for assays that can differentiate between acute infection, past exposure, and vaccination status. As Lapa et al. (2024) [1] emphasize, the clinical diagnosis of RVF is often challenging, particularly in the early stages, due to symptoms that overlap significantly with other viral hemorrhagic fevers (e.g., Ebola, Marburg, Crimean-Congo hemorrhagic fever) and common febrile illnesses (e.g., malaria, typhoid, dengue). This diagnostic ambiguity necessitates a dual-pronged approach, combining molecular detection of viral nucleic acids with serological identification of host antibodies. The World Health Organization (WHO) and the World Organisation for Animal Health (WOAH) recognize that no single validated point-of-care (POC) diagnostic tool currently exists for RVFV, underscoring the reliance on laboratory-based methods and the urgent need for field-deployable technologies [1, 41].

Molecular Diagnostics: Direct Pathogen Detection

Molecular methods are the primary tools for confirming acute RVFV infection, as they directly detect the viral genome, typically in blood or serum during the febrile viremic phase. The virus can only be reliably detected in the blood for a brief period, often 4-6 days post-onset of symptoms, making the timing of sample collection paramount [1]. Reverse transcription-polymerase chain reaction (RT-PCR) is the gold standard molecular technique, with real-time RT-PCR (RT-qPCR) offering high sensitivity, specificity, and quantitative capacity. These assays typically target conserved regions of the viral genome, most commonly the S-segment (encoding the nucleoprotein N) or the L-segment (encoding the RNA-dependent RNA polymerase) [5, 14]. The use of RT-qPCR has been instrumental in confirming outbreaks, as demonstrated during the 2016 outbreak in Uganda, where three of four acute human cases were confirmed by RT-PCR [37]. Similarly, during the 2015 outbreak in Mauritania, RT-PCR was used to confirm 57 of 184 suspected cases, enabling phylogenetic analysis that traced the virus to a Northeastern African lineage [46].

The utility of molecular diagnostics extends beyond human cases to animal surveillance and vector competence studies. For instance, RT-qPCR has been used to detect RVFV RNA in mosquito pools, confirming ongoing transmission during inter-epidemic periods (IEPs) in Senegal and Uganda [37, 39]. In livestock, RT-PCR has been critical for confirming acute infections in sheep, goats, and cattle, often before seroconversion occurs [48]. The development of high-throughput assays, such as the use of a recombinant RVFV MP-12 strain expressing Renilla luciferase, has facilitated the screening of antiviral compounds in a BSL-2 environment, bypassing the need for high-containment facilities for initial drug discovery [64].

However, molecular diagnostics have limitations. The short window of viremia means that a negative RT-PCR result does not rule out infection, particularly if the patient presents later in the disease course [1]. Furthermore, the requirement for sophisticated thermal cyclers, trained personnel, and a reliable cold chain for reagents and samples restricts the widespread deployment of RT-PCR in resource-limited, remote, or outbreak-prone settings. This has spurred the development of isothermal amplification techniques, such as loop-mediated isothermal amplification (LAMP), which can be performed at a constant temperature, reducing the need for expensive equipment. While promising, these assays have not yet been fully validated for field use in RVFV diagnostics [41].

Serological Diagnostics: Detecting the Host Response

Serological assays are indispensable for diagnosing RVFV infection beyond the viremic phase, for conducting seroprevalence surveys, and for confirming past exposure. The primary targets for serological detection are antibodies (IgM and IgG) directed against the viral nucleoprotein (N) and the envelope glycoproteins (Gn and Gc). The kinetics of the antibody response are well-characterized: IgM antibodies typically appear within 2-4 days post-onset of symptoms, peak around 7-10 days, and can persist for several months. IgG antibodies appear slightly later, around 5-7 days, and can persist for years, providing evidence of past infection [1, 5, 56].

Enzyme-Linked Immunosorbent Assays (ELISAs) are the most widely used serological platform for RVFV. Both indirect ELISAs (detecting total IgG or IgM) and competitive ELISAs (cELISAs, which detect antibodies that block binding of a labeled monoclonal antibody) are available. cELISAs are particularly valuable for screening across multiple species, as they are less dependent on species-specific secondary antibodies [28, 56, 62]. For example, a cELISA targeting the RVFV nucleoprotein has been used extensively in serosurveys of livestock, wildlife, and humans across Africa, revealing high seroprevalence rates in cattle (up to 34%) and goats (up to 32%) in hyperendemic areas of South Africa, even in the absence of reported outbreaks [55]. In Tunisia, a combination of ELISAs and indirect immunofluorescence assays (IIFA) confirmed a 2.3% seroprevalence of RVFV IgG in ruminants, providing the first serological evidence of viral circulation in the country [62].

Virus Neutralization Tests (VNTs) remain the gold standard for serological confirmation due to their high specificity. The plaque reduction neutralization test (PRNT) measures the ability of serum antibodies to neutralize live virus, preventing plaque formation in cell culture. PRNT is critical for differentiating RVFV from other phleboviruses and for confirming the specificity of ELISA-positive results [42, 55, 56]. However, VNTs require live virus, which must be handled in BSL-3 or BSL-4 facilities, limiting their use to specialized reference laboratories. To circumvent this, pseudovirus-based neutralization assays have been developed. These use replication-incompetent viruses (e.g., vesicular stomatitis virus pseudotypes) bearing the RVFV Gn and Gc glycoproteins, allowing neutralization testing under BSL-2 conditions [63]. This approach has been used to map neutralizing epitopes and evaluate vaccine-induced immunity.

The Imperative for Combined Molecular and Serological Approaches

The transient nature of RVFV viremia and the delayed onset of the antibody response create a diagnostic window where both molecular and serological tests may be negative. This is particularly problematic in the early febrile phase. Therefore, the most robust diagnostic strategy involves the simultaneous or sequential use of both RT-PCR and serology (IgM ELISA) on acute-phase samples, with a follow-up convalescent sample for seroconversion if the initial results are negative [1, 59]. This dual approach was critical in the 2016 Ugandan outbreak, where one of four cases was confirmed only by IgM and IgG serology, while the others were RT-PCR positive [37]. Similarly, in a study of febrile patients in Mozambique following severe flooding, 5% showed IgG seroconversion in convalescent samples, while all acute samples were RT-PCR negative, highlighting the role of serology in capturing cases missed by molecular testing [59].

The integration of diagnostics is also essential for surveillance. Seroprevalence studies using cELISA or PRNT provide a measure of past exposure and help define the geographic distribution and endemicity of RVFV [32, 49, 56]. For instance, a systematic review of seroprevalence studies from 1968 to 2016 found RVFV antibodies in 31 African countries, with significant variation by species and location [32]. In contrast, molecular surveillance using RT-PCR on mosquito pools or animal tissues can detect active viral circulation during IEPs, providing early warning of potential outbreaks [22, 44]. The combination of these approaches has revealed that RVFV is maintained through low-level cycling in livestock and wildlife, even during long inter-epidemic periods [22, 44, 60].

Emerging Diagnostic Technologies and Challenges

The development of POC diagnostics for RVFV remains a critical unmet need. Lateral flow assays (LFAs) for antigen or antibody detection are attractive for field deployment, but current prototypes lack the sensitivity and specificity required for reliable diagnosis [1]. The use of dried blood spots (DBS) for sample collection and transport has shown promise for serological and molecular testing, potentially expanding diagnostic access in remote areas [56].

Another frontier is the use of next-generation sequencing (NGS) and metagenomics for pathogen discovery and outbreak investigation. Whole-genome sequencing of RVFV from clinical samples, as performed during the 2016 Ugandan outbreak, allows for phylogenetic analysis, tracking of viral evolution, and identification of reassortment events [37, 40]. This is crucial for understanding the emergence of new lineages, such as the Northeastern African lineage responsible for the 2015 Mauritanian outbreak [46].

Despite these advances, significant challenges remain. The high genetic diversity of RVFV, driven by reassortment and mutation, can compromise the sensitivity of molecular assays if primer or probe binding sites are altered [47]. The lack of standardized, validated commercial assays across different laboratories hinders comparability of data. Furthermore, the cross-reactivity of serological assays with other phleboviruses (e.g., Toscana virus, Sandfly fever viruses) can lead to false positives, particularly in regions where these viruses co-circulate [25]. The development of a DIVA (Differentiating Infected from Vaccinated Animals) strategy is also critical for vaccination campaigns, requiring serological tests that can distinguish antibodies induced by natural infection from those elicited by vaccines lacking specific proteins (e.g., NSs) [17, 21, 65].

In conclusion, the advanced diagnostics for RVFV require a sophisticated, multi-layered approach. Molecular methods like RT-qPCR are essential for acute case confirmation and outbreak detection, while serological assays, particularly cELISA and PRNT, are indispensable for surveillance, retrospective diagnosis, and understanding the true burden of disease. The field is moving toward integrated diagnostic algorithms that combine these modalities, alongside the development of POC devices and genomic surveillance tools, to meet the challenges posed by this expanding zoonotic threat. The continued investment in diagnostic innovation is not merely a technical exercise but a fundamental pillar of global health security against emerging arboviruses.

Host Immune Responses and Vaccine Development Strategies

The intricate interplay between Rift Valley fever virus (RVFV) and the host immune system dictates the spectrum of clinical outcomes, ranging from subclinical infection to fatal hemorrhagic fever or encephalitis. A profound understanding of these immunological dynamics is paramount for the rational design of safe and efficacious vaccines, a critical need underscored by the WHO’s inclusion of RVFV on its Blueprint list of priority pathogens and the virus’s demonstrated capacity for geographical expansion into Europe, the Americas, and Asia via competent mosquito vectors [1, 4, 5, 31, 35, 41]. This section provides an exhaustive analysis of the host immune responses to RVFV infection and delineates the current landscape of vaccine development strategies, from traditional live-attenuated platforms to modern molecularly-defined candidates.

Innate Immune Evasion by the Viral NSs Protein

The cornerstone of RVFV pathogenesis is its capacity to subvert the host’s innate antiviral defenses, primarily orchestrated by the non-structural protein NSs. This major virulence factor acts as a pleiotropic antagonist of the type I interferon (IFN) system, which is the first line of defense against viral infection [3, 15]. Upon infection, host pattern recognition receptors (PRRs) such as RIG-I and MDA-5 detect viral RNA and signal through the adaptor protein MAVS (mitochondrial antiviral-signaling protein) to induce IFN-β production. However, NSs effectively dismantles this pathway through multiple mechanisms. It forms filamentous structures in the nucleus that sequester the transcription factor TFIIH subunit p62, leading to a global suppression of host cellular transcription, including the IFN-β gene [15]. Furthermore, NSs interacts with the SAP30-YY1 complex to bind chromatin and specifically repress IFN-β promoter activity [15]. The profound importance of MAVS in the central nervous system was demonstrated by Hum et al., who showed that Mavs-/- mice exhibit enhanced susceptibility to RVFV encephalitis, with increased brain viral burden, higher mortality, and dysregulated microglial responses, highlighting the non-redundant role of this pathway in neuroinvasion and control [7].

Beyond transcriptional suppression, NSs actively degrades key antiviral effector proteins. It targets protein kinase R (PKR), a critical sensor of double-stranded RNA that phosphorylates eIF2α to halt translation. Using a unique mechanism, NSs acts as a molecular bridge, recruiting PKR to the SCF (SKP1-CUL1-F-box) E3 ubiquitin ligase complex via the F-box proteins FBXW11 and β-TrCP1, thereby promoting PKR’s proteasomal degradation [18, 20]. This degradation is essential for maintaining viral protein synthesis and replication [18]. Interestingly, disrupting this SCF-NSs interaction with small molecule inhibitors or degron-mutant viruses blocks PKR degradation and leads to a paradoxical activation of PKR, which then potently suppresses viral replication, suggesting a therapeutic Achilles’ heel [18]. The structural complexity of NSs is further highlighted by its ability to form amyloid-like fibrils both in the nucleus and cytoplasm of infected cells, particularly in the brain of mice [8]. These fibrils, which exhibit typical properties of amyloids such as detergent resistance and Thioflavin-S binding, have been linked to increased mortality, adding a dimension of proteinopathy to viral pathogenesis [8]. The dual action of NSs, inhibiting IFN induction and degrading PKR, effectively paralyzes the innate immune system, allowing robust viral replication to proceed. This explains the exquisite susceptibility of most inbred mouse strains and many livestock species to wild-type RVFV, where infection often leads to overwhelming hepatic necrosis within days [48, 66].

Additional intrinsic cellular factors also shape the host-virus interface. The host RNA helicase DDX17 restricts RVFV replication by binding to and unwinding the virus’s non-coding RNAs, specifically the intergenic region (IGR) and 5’ non-coding region (NCR) of the S segment, thereby interfering with viral RNA synthesis [10]. Conversely, cellular pathways such as the Wnt/β-catenin signaling cascade are co-opted by the virus to create a favorable environment for replication; RNAi screening identified β-catenin as a host factor, and its inhibition reduces bunyavirus infection, suggesting a target for host-directed therapies [23]. The viral entry process itself is mediated by the Gn glycoprotein binding to the host receptor low-density lipoprotein receptor-related protein 1 (LRP1), and this interaction can be blocked by receptor-associated protein (RAP) domain 3, which protects mice from lethal challenge [2]. This discovery defines LRP1 as a critical entry factor and a promising target for antiviral intervention [2].

Adaptive Immunity: Humoral and Cellular Responses

While innate immunity is suppressed, the adaptive immune response is ultimately responsible for resolving infection and providing long-term protection. Both humoral (B cell) and cellular (T cell) arms contribute significantly, though neutralizing antibodies appear to be the primary correlate of protection [3, 11].

The humoral response is dominated by antibodies directed against the viral envelope glycoproteins, Gn and Gc. However, studies of naturally infected humans have revealed that the neutralizing antibody (nAb) response overwhelmingly targets Gn, rather than Gc [11]. Wright et al. demonstrated that while long-lived IgG (predominantly IgG1) and T cell responses are generated against both Gn and Gc, depletion of Gn-specific antibodies drastically reduces the neutralizing capacity of human sera, and IgG avidity for Gn, but not Gc, correlates with nAb titers [11]. This is consistent with the greater immunological accessibility of Gn on the virion surface. Structural and functional studies using human monoclonal antibodies (mAbs) have identified at least four major antigenic sites on the Gn-Gc heterodimer, with two key sites of vulnerability on Gn [6, 9]. Potently neutralizing mAbs, such as those isolated from MP-12 vaccinees or naturally infected survivors, have been shown to bind Gn domain A or complex quaternary epitopes that form only when Gn and Gc are co-expressed [6]. Importantly, many of these potent mAbs neutralize by a fusion inhibition mechanism; they bind to Gn and prevent the low-pH-induced conformational rearrangements in the Gn-Gc lattice that are required for Gc class II fusion loop exposure and membrane fusion [6, 9]. This mechanism is distinct from directly blocking the fusion loop. Representative mAbs targeting these sites have shown remarkable efficacy in mouse models, providing protection in both prophylactic and therapeutic settings, highlighting their potential as biologics for outbreak control [6, 12].

The cellular immune response also plays a vital role. CD4+ T helper cells are critical for providing help to B cells and for generating robust memory responses. CD8+ cytotoxic T lymphocytes (CTLs) can directly kill infected cells. In the ferret model of RVFV encephalitis, infection leads to lymphopenia early in disease, but animals that survive demonstrate cellular responses that likely contribute to viral clearance from peripheral tissues before neuroinvasion [53]. In mice, the NSs protein, by degrading PKR and inhibiting IFN signaling, not only blocks innate immunity but also impairs the development of a robust adaptive response. However, vaccines that lack NSs and thus allow for IFN induction tend to elicit stronger inflammatory and subsequently adaptive responses [3, 25].

Vaccine Development Strategies: From Bench to Bedside

Given the explosive nature of RVFV outbreaks, the lack of a licensed human vaccine, and the conditional use of veterinary vaccines, a massive global effort is underway to develop safe, effective, and DIVA (Differentiating Infected from Vaccinated Animals)-compatible vaccines for both livestock and humans. The strategies are diverse and exploit multiple platforms.

Live-Attenuated Vaccines (LAVs): Historically, LAVs have been the mainstay for veterinary use. The MP-12 strain, derived from a mutagenized virulent isolate, is conditionally licensed for livestock in the US and has been extensively tested in humans [41, 68]. Its attenuation is due to a combination of 23 mutations, including temperature-sensitive (ts) lesions in the L and M segments [68]. However, serial passage of MP-12 in cell culture can lead to reversion of certain ts mutations (e.g., L-G3750A), raising concerns about genetic stability and potential reversion to virulence, especially in immunosuppressed individuals [68]. Another prominent LAV, Clone 13, carries a large deletion in the NSs gene, rendering it highly attenuated and unable to suppress IFN [41]. While safe, Clone 13 can be less immunogenic, prompting efforts to create chimeric S segments that encode NSs from other phleboviruses (e.g., sandfly fever Sicilian virus) to improve immunogenicity while maintaining safety [25].

Reverse Genetics and Deletion Mutants: A significant advance has been the rational design of vaccines via reverse genetics. Deletion of the NSs gene alone (ΔNSs) or in combination with NSm (ΔNSs-ΔNSm) yields highly attenuated strains that are protective in multiple animal models, including non-human primates (common marmosets) [16]. These recombinant viruses induce robust nAb responses and protect against viremia and liver disease without causing clinical illness [16]. A particularly elegant innovation is the four-segmented RVFV (RVFV-4s) platform, where the M segment is split into two separate segments, each encoding one of the glycoproteins (Gn or Gc) [65]. This virus is genetically stable, does not cause encephalitis even when administered intranasally to mice, and, critically, is safe in pregnant ewes, the most sensitive target population, as it does not cross the placental barrier or cause fetal malformations or abortion [65]. This addresses a major safety hurdle for any RVFV vaccine.

Subunit and Virus-Like Particle (VLP) Vaccines: Recombinant protein subunit vaccines offer a high safety profile. The ectodomains of Gn and Gc, when expressed in eukaryotic systems, self-assemble into immunogenic complexes. A subunit vaccine based on the Gn and Gc glycoproteins has been shown to elicit high-titer nAbs in sheep and confer complete protection against heterologous virulent challenge (Kenya-128B-15), with no viremia, fever, or histopathological lesions observed post-challenge [21]. This platform is inherently DIVA-compatible, as vaccinated animals lack antibodies to the non-structural proteins (NSs, NSm) that are produced during natural infection [21]. DNA vaccines encoding the Gn ectodomain have also been tested but generally induce lower, non-neutralizing antibody titers, though they can still confer partial protection in sheep, correlating with anti-eGn IgG levels rather than T cell responses [17].

Viral Vector Vaccines: Recombinant viral vectors represent another promising avenue. Vesicular stomatitis virus (VSV) pseudotypes bearing RVFV Gn and Gc can be used for safe in vitro nAb assays under BSL-2 conditions and also serve as immunogens in vivo [63]. Similarly, a recombinant rabies virus (RABV) vector expressing the RVFV Gn ectodomain (rSRV9-eGn) induced RVFV-specific IgG and T cell responses, albeit without detectable nAbs, in mice [67]. While the lack of nAbs is a limitation, such a bivalent vaccine could address the dual burden of RVF and rabies in Africa [67].

Correlates of Protection and Considerations for Human Vaccination

The convergence of evidence from natural infection and vaccination studies strongly indicates that neutralizing antibodies targeting the Gn glycoprotein are the primary correlate of protection against RVFV [6, 11, 21]. However, T cell responses, while not sufficient alone, likely contribute to viral clearance and long-term memory [3, 67]. For human vaccine trials, which are now being actively planned given the expanding epidemiology of RVFV [4], several key considerations are paramount. First, safety in pregnant women and immunocompromised individuals is non-negotiable, favoring platforms like subunit, VLP, or rationally attenuated LAVs such as RVFV-4s over traditional LAVs [65]. Second, the ability to rapidly scale production during an outbreak is critical, favoring platforms with established manufacturing pipelines (e.g., subunit, virus-like particles). Third, a DIVA strategy will be essential for veterinary vaccines to allow serological surveillance and trade. The successful use of multiple platforms in target species, sheep, cattle, non-human primates, provides a clear regulatory pathway, making an effective human RVFV vaccine an achievable goal in the near future.

Prevention, Surveillance, and One Health Control Measures

The prevention and control of Rift Valley fever virus (RVFV) necessitates a paradigm shift from reactive outbreak response to proactive, integrated strategies that embrace the interconnectedness of human, animal, and environmental health. This One Health approach, championed by the World Health Organization (WHO), the World Organisation for Animal Health (WOAH), and the Food and Agriculture Organization (FAO), is not merely an aspirational framework but an operational imperative given the virus's complex ecology, vector-borne transmission, and zoonotic potential. The pathogen’s capacity for rapid emergence, its maintenance during prolonged inter-epidemic periods (IEPs), and the expanding geographic range of competent vectors demand a multi-pronged strategy that integrates advanced vaccination platforms, robust surveillance networks, and ecologically-informed vector management.

Vaccination Strategies: The Cornerstone of Prevention

Vaccination remains the single most effective intervention for mitigating the devastating impact of RVFV in both livestock and humans. The development of safe, efficacious, and differentiating infected from vaccinated animals (DIVA)-compatible vaccines is a critical priority. Several promising platforms are under investigation, each with distinct advantages and limitations.

Live-Attenuated Vaccines: The MP-12 vaccine, a conditionally licensed live-attenuated strain for veterinary use in the United States, has demonstrated immunogenicity in livestock and humans [68]. However, its genetic stability is a concern. Serial passage studies have identified reversion mutations at specific loci, including the temperature-sensitive mutation L-G3750A, highlighting the necessity of a stringent seed lot system to prevent reversion to virulence [68]. The NSs deletion mutant (ΔNSs) and double deletion mutant (ΔNSs-ΔNSm) represent safer, rationally designed alternatives. These reverse-genetics-derived candidates are highly attenuated and protective in non-human primates and sheep, lacking the NSs virulence factor responsible for interferon antagonism [16]. A critical breakthrough in safety is the development of four-segmented RVFV (RVFV-4s) variants. By splitting the M segment, this virus cannot reassort with wild-type strains and, crucially, does not cross the placental barrier in pregnant ewes, preventing the teratogenic effects and abortion storms characteristic of natural infection [65]. This represents a monumental advance for use in pregnant livestock, a previously prohibitive population for vaccination.

Subunit and DNA Vaccines: Recombinant subunit vaccines based on the Gn and Gc glycoproteins offer a DIVA-compatible solution. A prototype vaccine formulated with these proteins conferred complete protection in sheep against heterologous challenge, preventing viremia, fever, and histopathological lesions [21]. DNA vaccines encoding the Gn ectodomain have shown partial protection in sheep, with protection correlating with anti-Gn IgG responses rather than robust T-cell responses [17]. While these platforms are exceptionally safe, they often require multiple doses or potent adjuvants to achieve durable, neutralizing antibody titers comparable to live-attenuated vaccines.

Novel Platforms and Passive Immunization: Chimeric vaccines utilizing viral vectors, such as a recombinant rabies virus expressing Gn, induce humoral and cellular immunity in mice, offering the dual benefit of protecting against both RVFV and rabies [67]. Pseudovirus-based systems allow for the safe, high-throughput screening of neutralizing antibodies under BSL-2 conditions, accelerating vaccine evaluation [63]. Furthermore, human monoclonal antibodies (mAbs) isolated from survivors and vaccinees demonstrate extraordinary potency. mAbs targeting the Gn domain or the Gc/Gn hetero-oligomer inhibit viral fusion at picomolar concentrations and protect mice in both prophylactic and therapeutic settings [6]. This passive immunization strategy provides an immediate countermeasure for high-risk exposures, such as laboratory accidents or outbreaks in vulnerable populations, and defines critical sites of vulnerability on the viral surface for structure-based vaccine design [12].

Surveillance: The Sentinel Net for Early Detection

Effective surveillance is the nervous system of any RVFV control program, providing the early warning necessary to trigger preventive actions. The challenge is that the virus circulates silently during IEPs, making passive case reporting grossly inadequate.

Serological Surveillance and Inter-Epidemic Circulation: Decades of research have dismantled the myth that RVFV is entirely quiescent between outbreaks. Serological surveys in livestock have consistently demonstrated ongoing, low-level transmission. In South Africa, seroconversion rates in cattle and goats reached 0.59 and 0.41 per animal-year, respectively, in the absence of reported outbreaks, indicating a hyperendemic state [55]. Similar findings in Mozambique (44.2% seroprevalence in sheep), Tanzania (8.2%), and Kenya confirm that the virus is maintained through cryptic cycles involving livestock, wildlife, and possibly humans [44, 49, 60]. Camels also play a significant role, with seroprevalence rates of 14.2% in Kenya, often in mixed infections with Brucella spp. and Coxiella burnetii, complicating diagnosis [28]. Critically, infection during pregnancy is strongly associated with miscarriage in humans, as demonstrated in a Sudanese cohort where 54% of RVFV-infected pregnant women experienced fetal loss [38]. This underscores that surveillance must extend beyond animal health to include reproductive health outcomes in endemic regions.

Entomological Surveillance and Vector Ecology: Mosquito surveillance must be integrated and predictive. The virus is maintained transovarially in Aedes mosquitoes, allowing infected eggs to survive desiccation for years, serving as the primary mechanism for outbreak initiation after heavy rainfall [27]. Culex species then amplify the virus among livestock. Vector competence studies have identified numerous species capable of transmission, including Culex pipiens and Aedes albopictus in Europe, and Aedes vexans and Culex tritaeniorhynchus in Senegal, whose abundance is driven by rainfall, temperature, and specific biotopes [31, 35, 39]. Remote sensing data, such as satellite-derived rainfall and temperature estimates, can be integrated into population dynamics models to forecast vector abundance and create dynamic risk maps, providing a powerful tool for targeted vector control interventions weeks before an outbreak becomes clinically apparent [43].

Genomic and Environmental Surveillance: Phylogenetic analyses have revealed the dynamic phylogeography of RVFV, with Kenya acting as a major source population for introductions into Madagascar, Egypt, and the Arabian Peninsula [40]. The identification of a Northeastern African lineage during the 2015 Mauritanian outbreak highlights the ongoing evolution and transboundary spread of the virus [46]. Surveillance systems must therefore include real-time genomic sequencing to track viral lineages, detect reassortment events (which have been implicated in increasing human virulence), and monitor for the emergence of vaccine escape mutants [47]. Wastewater surveillance, while not yet validated for RVFV, represents a potential future tool for population-level monitoring in both livestock and human communities.

One Health Control Measures: Bridging Sectors

The operationalization of a One Health approach requires coordinated action across veterinary, public health, and environmental sectors. Control measures must be tailored to the epidemiological context, whether that involves preventing an incursion into a naïve region or mitigating an ongoing epizootic.

Livestock Movement and Trade Restrictions: The trade of live animals is a primary driver of long-distance viral spread. The 2000 outbreak in Saudi Arabia and Yemen was directly linked to the importation of infected livestock from the Horn of Africa [5]. Therefore, pre-movement testing, quarantine protocols, and trade bans from endemic zones during active outbreaks are essential. However, modeling studies from the Comoros archipelago suggest that coordinated within-island control (e.g., vaccination) is more effective than between-island movement restrictions alone [34]. This highlights that trade bans must be coupled with robust local vaccination and vector control.

Vector Control and Environmental Management: Targeted vector control can reduce transmission, but it must be ecologically intelligent. Larviciding of Aedes breeding sites (temporary ponds, irrigation canals) before the rainy season can reduce the primary inoculum. Adulticiding with pyrethroids can be used during outbreaks, but insecticide resistance must be monitored. The use of Wolbachia-infected mosquitoes, which can block pathogen transmission, is a promising biocontrol strategy, though its efficacy is pathogen-specific. For RVFV, Wolbachia (wAlbB) showed no significant effect on infection or transmission in Culex tarsalis, contrasting with its strong blocking of dengue in Aedes aegypti [69]. This emphasizes that novel control tools must be rigorously tested against the target pathogen in the relevant vector species. Environmental modifications, such as improved water management in irrigation schemes, which have been shown to increase seroconversion risk [44], can reduce mosquito breeding habitat.

Community Engagement and Risk Communication: Ultimately, control measures fail without community buy-in. Studies from Tanzania reveal that livestock keepers often do not perceive RVF as a priority disease, despite recurrent outbreaks, leaving them poorly prepared [60]. Public health messaging must address specific risk behaviors, including the consumption of raw milk, which is a significant risk factor for human infection, with seropositive individuals 2.5 times more likely to consume raw milk [36]. Messaging must also emphasize the risks associated with handling aborted animal material and unprotected contact with sick animals [4, 61]. For pregnant women, the documented link between RVFV infection and miscarriage necessitates targeted counseling during outbreaks [38, 54]. A fully integrated One Health surveillance system, one that links veterinary data on abortion storms and seroconversions with human febrile illness surveillance, mosquito abundance indices, and real-time climate data, provides the only framework capable of predicting, preventing, and controlling this formidable pathogen.

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