Translational Model of Zika Virus Disease in Baboons

KEYWORDS: baboon, West Nile virus, ZIKV, Zika virus, flavivirus, nonhuman primate

INTRODUCTION

Zika virus (ZIKV) was initially isolated from a febrile sentinel rhesus monkey in the Zika Forest in Uganda in 1947 (1) and belongs to the family Flaviviridae, genus Flavivirus, which includes dengue (DENV), West Nile (WNV), yellow fever (YFV), and Japanese encephalitis (JEV) viruses (2). While Aedes sp. mosquitoes are the primary vectors for ZIKV (Aedes aegypti) (3), recent outbreaks have shown that the virus can be transmitted vertically, sexually, and through blood transfusions (4,–6). Prior to the 2007 ZIKV outbreak on Yap island in the Federated States of Micronesia (∼5,000 cases), there had been only 14 documented cases of human ZIKV infection, restricted to Africa and Southeast Asia. The subsequent French Polynesian ZIKV epidemic (2013–2014) resulted in infection of ∼11% of the population (∼30,000 cases) (7). Although ZIKV infection was originally viewed as self-limiting, relatively harmless, and nonlethal, with symptoms of mild fever, rash, conjunctivitis, arthritis, and arthralgia (8), the South Pacific epidemics marked a notable shift of ZIKV infection with increased severity, including neurological complications, such as Guillain-Barre syndrome (9). Retrospectively, an unusual increase in fetal central nervous system (CNS) malformations, including microcephaly, was associated with the French Polynesian ZIKV outbreak. Furthermore, the ZIKV epidemic in Latin America (more than 1.5 million cases in Brazil since 2015) confirmed the link between ZIKV and fetal microcephaly, as well as other fetal brain malformations, including cerebral calcifications, cerebral atrophy, ventriculomegaly, parenchymal brain hemorrhages, and reduced middle cerebral artery blood flow, and other fetal anomalies, including intrauterine growth restriction (IUGR) (4, 9).

It is critical to establish animal models with translational significance for humans in order to study the pathogenesis of ZIKV infection and to provide accurate models for testing novel interventions, including vaccines and antiviral compounds. Numerous studies have employed mouse models for studying ZIKV infection. These have largely relied upon transgenic mice either lacking the ability to produce or respond to interferons (IFNs) (10, 11) or passively immunoneutralized against IFNs (11, 12). There are limited reports where wild-type mice have been used to study ZIKV infection. However, in these studies, the virus was delivered either directly to the immune-incompetent fetus in utero or directly into neonatal pups. Direct inoculation of ZIKV into fetuses or neonates also does not recapitulate the human pregnancy paradigm, since the noted placental pathologies leading to placental compromise in ZIKV-infected human pregnancies likely contribute to the fetal outcome, including IUGR and CNS malformations (4, 13, 14).

Nonhuman primates (NHPs) are the best-documented animal reservoirs for ZIKV (and related flaviviruses), and ZIKV infection has been studied in rhesus (Macacca mulatta) (15,–24), cynomolgus (Macacca fascicularis) (17, 25), and pigtail (Macacca nemestrina) (26) macaques, using males and nonpregnant and pregnant females (15,–17). In several studies of rhesus and cynomolgus macaques, subcutaneous inoculation with the H/PF/2013 (French Polynesia, 2013) (15), PRVABC59 (Puerto Rico, 2015) (16, 17, 25), Brazilian (Brazil/ZKV2015; Zika virus/H.sapiens-tc/BRA/2015/Brazil_SPH2015) (21,–24), or Cambodian (FSS13025) (25) ZIKV strain led to systemic viremia in both males and nonpregnant females that peaked between 2 and 6 days postinfection (dpi) and typically resolved by 5 to 10 dpi in males and nonpregnant females. ZIKV RNA has been detected in saliva, urine, cerebrospinal fluid (CSF), spinal cord, brain, peripheral nervous tissue, lymph nodes, semen, uterus, and vagina, albeit transiently. There was variability in both the duration and presence of the virus in the various body components between the studies. Notably, ZIKV RNA was detected in neuronal, lymphoid, saliva, and joint/muscle tissues and male and female reproductive tissues several weeks postinfection (16, 22, 23). A robust immune response correlated with the resolution of systemic viremia was observed and provided protective immunity upon reinfection with either a homologous or heterologous ZIKV strain (17). One study that evaluated the systemic inflammatory response following subcutaneous inoculation (16) noted a weak cytokine response. Most, but not all, studies reported that ZIKV was accompanied by transient fever, rash on the arms and torso, lymphadenopathy, and conjunctivitis (15,–17). ZIKV infection has also been achieved in macaques via an oral mucosal route of administration (20) and vaginal delivery of the virus (21). ZIKV infection has been reported in pregnant macaques, as well (15, 18, 27,–29). Of interest, the major observation in the infected pregnant rhesus macaque was prolonged viremia lasting approximately 1 month or more, despite developing neutralizing antibodies. New World primates, such as the owl monkey (Aotius sp.), squirrel monkey (Samiri sp.), and marmoset (30, 31) have also been reported as models of ZIKV. In these models, viremia, as well as the presence of virus in other body fluids, such as semen and saliva, were detected, and seroconversion was reported. However, there was no evidence of clinical disease.

In the present study, we developed the olive baboon (Papio anubis; adult males and nonpregnant females) as an alternative NHP model to study ZIKV infection and pathogenicity that can be correlated with the human situation and compared/contrasted with ZIKV infection in other nonhuman primates. The olive baboon is similar to humans in terms of size, genetics, reproduction, placentation, brain development, and immune repertoire (all four subclasses of IgG), which makes the baboon an excellent translational model to study ZIKV infection and for vaccine and therapeutics development (32,–34). Studies have utilized the baboon as an NHP model for assessing the safety and efficacy of vaccines in adults and in pregnant females and their infants (33, 35). The baboon is also permissive to flavivirus infection and replication and produces a virus-specific immune response (34, 36). Here, we describe infection of olive baboons with a contemporary French Polynesian strain of ZIKV (H/PF/2013) that is associated with the emergence of neurological disorders in adults and congenital Zika syndrome in the Americas (37,–39). It was recently shown that a single point mutation in the prM protein of ZIKV in the Asian strain arose before the 2013 outbreak in French Polynesia and that it results in increased ZIKV infectivity in both human and mouse neuroprogenitor cells (40).

Go to:

RESULTS

Description of animal cohorts and experimental outline.

For this study, adult male (n = 3) (M1, M2, and M3) and female (n = 3) (F1, F2, and F3) baboons were used. The baboons were either infected with a single dose of 1 × 10⁶ focus-forming units (FFU) ZIKV (H/PF/2013; n = 3 male baboons) or 1 × 10⁴ FFU (n = 3 female baboons). Anesthetized animals were inoculated with a single clinically relevant dose of the virus administered subcutaneously in the midscapular area. Blood, urine, saliva, and CSF were collected at preinfection (day 0) and subsequent days postinfection as shown in Fig. 1. The baboons were euthanized at 41 (male) and 43 (female) days postinfection. All the animals developed a rash at the site of inoculation, on the abdomen, and in the inguinal and axillary regions. In addition, all the baboons developed conjunctivitis. None of the animals showed signs of any clinical disease, such as weight loss, other than those described above.

 

 

 

FIG 1

Timeline of ZIKV infection and sample collection from male and female olive baboons. Adult nonpregnant female and male baboons (n = 3) were infected subcutaneously with ZIKV at 10⁴ FFU and 10⁶ FFU, respectively. For the female study, blood and saliva were collected at 0, 2, 4, 8, 14, 21, and 43 dpi. Urine was collected at 0, 2, 4, 6, 10, 14, 21, and 43 dpi and CSF at 0, 2, 8, 14, 21, and 43 dpi. For the male study, blood was collected at 0, 3, 4, 5, 6, 7, 11, 13, 20, 27, 34, and 41 dpi. Urine and saliva were collected at 0, 4, 6, 11, 13, 20, 27, 34, and 41 dpi and CSF at 4, 6, 11, 13, 20, 34, and 41 dpi. Necropsies were performed at 43 dpi for the female study and 41 dpi for the male study.

Complete blood counts (CBCs) were evaluated for all the males and females on EDTA-anticoagulated whole-blood samples collected on day 0 and subsequent days postinfection, as shown in the experimental timeline (Idexx ProCyte DX hematology analyzer; Idexx Laboratories, ME). The CBCs included analysis for red blood cells (RBCs), hemoglobin, hematocrit, and platelet count. RBC, hemoglobin, and hematocrit numbers did not show any differences pre- and post-ZIKV infection in all the males and females. Notably, platelet numbers went down in 2 out of 3 male baboons (M1 and M2) in the acute phase of the infection (5 dpi), followed by an elevation in platelet numbers in all the male baboons by day 15 postinfection and return to preinfection numbers by day 41 postinfection (Fig. 2B). In the female cohort, one baboon had a noted decline in platelets from days 6 to 14 postinfection and a second exhibited a mild delayed drop in platelets by day 22 postinfection, while platelet counts in the third were unaffected by ZIKV infection (Fig. 2A). Although elevation and decline in platelet numbers were observed in some baboons (males and females), the numbers were within the normal ranges for platelets for male and female baboons (41).

 

FIG 2

Whole-blood platelet counts from male and female baboons. CBCs were performed on EDTA-anticoagulated whole blood for female (A) and male (B) baboons. Platelet counts per milliliter of whole blood are shown for the specified time points postinfection with ZIKV.

Viral load data postinfection in whole blood.

Viral RNA (vRNA) was quantified by one-step quantitative reverse transcription (qRT)-PCR in RNA extracted from the blood samples. Blood samples were collected from female baboons on day 0 and days 2, 4, 8, 14, 21, and 43 postinfection. ZIKV RNA was detected in the blood of one baboon (F3) on day 2 postinfection and in the other two by day 4 postinfection. Peak viremia occurred on day 4 postinfection in all three low-inoculum-titer baboons (range, 2.2 × 10⁴ to 5.3 × 10⁴ copies/ml) and was undetectable by day 14 postinfection (Fig. 3A). Blood samples from male baboons were collected preinfection on day 0 and 3, 4, 5, 6, 7, 11, 13, 20, 27, 34, and 41 days postinfection. Viremia was detected in all 3 baboons on day 3 postinfection. Peak viremia occurred between days 3 (M2 and M3) and 4 (M1) postinfection (range, 7.9 × 10³ to 4.0 × 10⁵ copies/ml) and was undetectable by day 11 postinfection (Fig. 3B).

 

 

Open in a separate window

FIG 3

ZIKV loads in blood, mucosal fluids, and CSF from infected male and female baboons. ZIKV RNA was extracted from specimens collected from each animal at the indicated days postinfection and quantitated by one-step qRT-PCR in whole blood, saliva, CSF, and urine from females (A, C, E, and G) and males (B, D, F, and H).

ZIKV shedding into mucosal fluids (saliva and urine) and CSF.

Saliva and urine samples from the female cohort were collected at the same frequency as the blood samples. ZIKV RNA was undetectable in the urine samples from all three baboons (Fig. 3C). In saliva, ZIKV RNA was detectable on day 8 following infection in two out of three low-titer-inoculum baboons (F1 and F2), and in one animal (F2), ZIKV RNA was detectable until day 10 postinfection, longer than blood viremia detection. Peak viremia in saliva was 8.3 × 10⁴ copies/ml. Baboon F3 did not have detectable ZIKV RNA in saliva even though viral RNA was detected in her blood (Fig. 3E). CSF was collected from the cisterna magna in the cohort on days 0, 2, 8, 14, 21, and 43 after infection. ZIKV RNA was detected in the CSF of one baboon (F2) on day 8 (Fig. 2G) postinfection.

In the male cohort, saliva and urine samples were collected on days 4, 6, 11, 13, 20, 27, 34, and 41 postinfection. Viruria was detected in one out of three baboons (M1) and was first detected on day 6 and persisted until day 27 (Fig. 3D), similar to what was observed for saliva in the animal. The peak viruria in this animal was on day 11 postinfection (1.4 × 10⁶ copies/ml). The other 2 males did not have detectable ZIKV RNA in the urine (Fig. 3D). Two out of three baboons had detectable viral RNA in the saliva (Fig. 3F). In one (M1), viral RNA was detected in saliva on day 6 and persisted until day 27 postinfection, again, longer than detection in the blood. The other baboon (M2) had low viral RNA copy numbers on day 4 (1.3 × 10³ copies/ml), and they were undetectable in subsequent saliva samples. M3 did not have viral RNA in its saliva despite detectable blood viremia (Fig. 3D).

CSF was collected from the cisterna magna in this cohort on days 4, 6, 11, 20, 34, and 41 postinfection. ZIKV RNA was detected as early as day 4 in one baboon (M3) and on day 11 in another (M1) (Fig. 3H). ZIKV RNA was detected in all the collected samples, which included blood; mucosal fluids, such as urine and saliva; and CSF in M1, with the highest peak viremia in the blood.

ZIKV RNA in tissue samples obtained at necropsy.

Reproductive tissues (testes, epididymis, seminal vesicles, and prostate), bladder, and lymph nodes (axial, inguinal, mesentery, and submandibular) were collected from the male baboons during necropsy on day 41 postinfection. ZIKV RNA was detected in the left epididymis of M1 (3 × 10⁴ copies/mg). Apart from this one tissue from M1, viral RNA was undetectable in any of the reproductive tissues in all the males 41 days postinfection. However, vRNA was detected in various lymph nodes in all three male baboons at necropsy. M1 had vRNA in the submandibular lymph nodes (3.6 × 10⁴ copies/mg), M2 had vRNA in the mesenteric lymph nodes (1.6 × 10³ copies/mg) and inguinal lymph nodes (9.8 × 10⁴ copies/mg), and M3 also had ZIKV RNA in the mesenteric lymph nodes (1.2 × 10⁴ copies/mg) and inguinal lymph nodes (2.0 × 10⁴ copies/mg) (Fig. 4). No vRNA was detected in tissues collected from female baboons at day 43 postinfection, including ovarian, uterus, and vaginal epithelium tissues. Lymph nodes were not collected from this group at the time of the necropsy.

 

 

FIG 4

ZIKV loads in tissues collected at necropsy. ZIKV RNA was extracted from flash-frozen tissue samples collected at necropsy on day 41 postinfection and quantitated by one-step qRT-PCR. LT, left; RT, right; Sem. Ves., seminal vesicle; Ing. LN, inguinal lymph node; Mes LN, mesenteric lymph node.

ZIKV antibody response.

ZIKV-specific IgM and IgG responses in the sera of female and male baboons were quantified by enzyme-linked immunosorbent assay (ELISA). Anti-IgM antibodies were detectable in sera collected from female baboons between days 10 and 40 postinfection, with a peak on day 14 (Fig. 5A). One animal (F3) peaked at day 10 and showed reduced values compared to the other low-titer-inoculum baboons, and by day 30 postinfection, IgM levels were below the positive cutoff value for the animal. Baboon F2 demonstrated a robust IgM response, with absorbance values for ZIKV IgM antibody four times the value of the positive-control serum, which remained elevated compared to other subjects in the cohort until day 40 postinfection. Anti-ZIKV IgG was detectable in F2 by day 14 postinfection and in the remaining two baboons by day 28 (Fig. 5C). Similar to the IgM response, animal F2 had a robust ZIKV IgG response, with absorbance values for ZIKV IgG twice the value of the positive-control serum on day 14 and remaining elevated compared to the remainder of the cohort on day 28 postinfection.

 

 

FIG 5

Anti-ZIKV IgM and IgG and anti-WNV IgG responses in male and female baboon sera. Sera from infected baboons were tested by ELISA for antibodies (Abs) directed against ZIKV infection. IgM (A) and IgG (C) antibodies in female sera and IgM (B) and IgG (D) antibodies in male sera were tested at the indicated days postinfection. Male (F) and female (E) sera from ZIKV-infected baboons were also tested by ELISA for the presence of IgG antibodies against WNV. Female F2 and male M3 tested positive for WNV infection. The dashed lines represent the assay cutoff controls for IgM and IgG detection in the samples.

In sera from the male baboons, Anti-ZIKV IgM antibody was detected in one baboon, M1, by day 8 postinfection and in all the animals until day 34. Animal M1 showed a robust IgM response to ZIKV, with 8 times the absorbance value of the positive control on day 14 and higher than the rest of the baboons in the cohort, with levels remaining elevated through day 34 postinfection (Fig. 5B). A ZIKV-specific IgG response was detectable in 2 of the 3 baboons by day 14 postinfection and was detectable in all the animals at day 28 (Fig. 5D). The IgG response to ZIKV was elevated in M1 at day 28 postinfection, similar to its elevated IgM levels compared with the remainder of the cohort.

In addition, neutralizing antibodies were determined for each baboon on day 28 postinfection using a plaque reduction neutralization test (PRNT). All the baboons infected with ZIKV developed neutralizing titers by day 28 postinfection (PRNT₅₀ [the concentration of serum required to neutralize 50% of the plaque count of a known amount of serum-free ZIKV] endpoint titers: females [10⁴-FFU inoculum], 1:1,280 [F1], 1:1,280 [F2], and 1:640 [F3]; males [10⁶-FFU inoculum], 1:640 [M1], 1:1,280 [M2], and 1:640 [M3]). Furthermore, endpoint dilution titers were very consistent among the cohorts, with no significant differences. Preinfection sera did not demonstrate any ability to neutralize ZIKV.

WNV antibody response.

WNV IgG was detected in the sera from baboon F2 on days 0, 8, 14, 30, and 43, with the peak response on day 14 (Fig. 5E), and in baboon M3 on days 0, 7, 13, 27, and 41, with the peak response on day 27 (Fig. 5F). The other two baboons in each cohort were not seropositive for WNV and did not elicit an anti-WNV IgG response following ZIKV inoculation.

Cytokine response to ZIKV infection.

Male and female baboons had distinct cytokine responses to ZIKV infection. There was variability in the plasma levels of the different cytokines between the two cohorts and between the acute (3 or 4 to 14 dpi) and convalescent/latent (>14 dpi until termination) phases postinfection. Despite being inoculated with a 100-fold lower ZIKV load, female baboons tended to have a more pronounced acute cytokine response than the male baboons (10⁶-FFU inoculum). In general, the male baboons exhibited a more latent cytokine response (convalescent phase) to ZIKV infection than the female baboons. In particular, in the female cohort, a significant (P ≤ 0.05) increase in the acute phase (≤10 dpi) was noted for monocyte chemoattractant protein 1 (MCP-1), interleukin 2 (IL-2), IL-8, IL-10, and IL-15 (Fig. 6A). A trend (P ≤ 0.08) toward elevated IFN-γ (P = 0.08) and IL-12 (P = 0.08) was also noted. In addition, in this cohort, we observed a significant (P < 0.05) decrease in plasma soluble CD40 ligand (SCD40L) in the acute phase. In female baboons in the convalescent phase, we observed significant (P < 0.05) elevations in plasma MCP-1, IL-8, and IL-2, with trends toward elevated IL-6 (P = 0.08) and transforming growth factor α (TGF-α) (P = 0.08).

 

 

FIG 6

Cytokines and growth factor expression in baboon sera following ZIKV infection. Cytokine and growth factor expression for the female (A) and male (B) baboons were determined during the acute (bars A) and convalescent/latent (bars L) phases post-ZIKV infection. A paired t test was used to test for differences between preinfection (day 0) plasma cytokine concentrations and peak plasma cytokine concentrations obtained in the acute phase (≤10 dpi) and the convalescent phase (≥14 dpi) for each baboon, since individual starting concentrations were highly variable between animals, as was the day of the peak plasma cytokine response. All the data are presented as means and standard errors of the mean (SEM). The numbers above the bars are P values; bars without the numbers have P values of >0.1.

In the male baboons, significant increases in plasma cytokine levels in the acute phase were more restricted, with increases (P < 0.05) noted for MCP-1 and IL-8, with a trend toward increased IL-15 (P = 0.08), while SCD40L was significantly decreased (Fig. 6B). In the convalescent phase, significant increases were noted for IL-8, granulocyte colony-stimulating factor (G-CSF), TGF-α, and SCD40L.

ZIKV cellular immunity.

Peripheral blood mononuclear cells (PBMCs) were assayed by cytokine enzyme-linked immunosorbent spot (ELISPOT) assay for the number of cells expressing IFN-γ in response to stimulation with either ZIKV envelope protein or NS1. As a positive control, matching aliquots of all samples were stimulated with phytohemagglutinin (PHA). As expected, following stimulation with the strong mitogen PHA, all the samples were highly positive for IFN-γ expression. PBMC aliquots from both cohorts demonstrated antigen-specific production of IFN-γ following stimulation with both envelope and NS1 proteins (Fig. 7).

 

 

FIG 7

ELISPOT assay for detecting ZIKV antigen-specific IFN-γ-producing cells. Aliquots of PBMCs from ZIKV-infected baboons were stimulated with either ZIKV envelope protein, ZIKV NS1 protein, or PHA as a positive control. The total number of SFCs in duplicate wells was determined for each sample and adjusted to that for negative-control medium-only background wells.

Go to:

DISCUSSION

In this study, we describe Zika virus infection of the olive baboon (Papio anubis) as a novel nonhuman primate model for the translational study of ZIKV. Nonhuman primate models of flavivirus infection, specifically Zika virus, offer unique opportunities to study the biology and pathogenesis in a translational setting. Established models such as the baboon allow the fulfillment of a necessary prerequisite for preclinical testing of novel vaccines and therapeutics prior to conducting clinical trials. While other animal models of Zika virus have been developed, including mice and other nonhuman primates (e.g., macaques), the olive baboon provides an additional translational model that closely resembles the disease state observed in humans (Table 1). Mice offer an excellent model for analyzing the principal components of host-pathogen interactions, and the seemingly endless genetic models provide numerous prospects for studying those interactions. However, immunocompetent mice are refractory to infection with Zika virus unless some intervention is used to block the type 1 interferon pathway or other immunodeficiencies are induced by genetic means, raising questions about the translational reliability of these findings from mice to humans (11, 12).

TABLE 1

Comparison of cytokine expression levels^a

 

^aNC, no change; LD, below the limit of detection; NR, not reported; L, latent phase (convalescent); A, acute phase; T, trend (P > 0.05); T*, increase reported but no statistical analysis provided. Data with a P value of <0.05 are shaded.

^bReference 1.

^cReferences 16 and 22.

Other established nonhuman primate models for the study of Zika virus are rhesus, cynomolgus, and pigtail macaques (15, 16, 26), as well as New World primates, such as the owl monkey (Aotius sp.), squirrel monkey (Samiri sp.), and marmoset (30, 31). While multiple studies have reported clinical disease in the rhesus model that is similar to that observed in humans, others have found limited clinical symptoms (42). One possibility to explain the reported differences could be the different origins of the macaque species, e.g., Indian- versus Chinese-derived rhesus macaques, as animals from these two origins are known to have genetic differences (43). Cynomolgus monkeys inoculated with Zika virus demonstrate viral replication in many tissues yet fail to demonstrate clinical symptoms, as well (25). A study of ZIKV infection in a single pregnant pigtail macaque at approximately three-fourths of the gestation period found significant fetal brain pathologies. However, the very high inoculation titer used in that study (five subcutaneous inoculations of 10⁷ PFU each), coupled with the use of a Cambodian strain of ZIKV not associated with microcephaly or neuropathologies in humans, raises questions about the translational significance of the findings. Interestingly, this animal also did not present the symptoms associated with ZIKV infection, i.e., rash, conjunctivitis, or fever (26). The presentation of clinical symptoms after Zika virus infection is important in the context of pregnancy, since studies done in pregnant Brazilian women infected with Zika virus have shown that one in five of all pregnant women with Zika virus infection present a rash. Adverse fetal outcomes were also more common in Zika virus-positive pregnant women with clinical symptoms than in those without symptoms (4, 14). Taking these observations into consideration, not a single model reported to date stands out as a gold standard for translational studies. As such, it is important to continue to study other species of nonhuman primates used in research to determine their usefulness as a translational model of Zika virus. Furthermore, contrasting and comparing the differences in responses to ZIKV that may occur between macaques and baboons should provide invaluable insight into pathologies associated with ZIKV, such as Guillain-Barre syndrome, other neuropathologies, and congenital pathologies in pregnancy.

Baboons have been used to study human infectious diseases and to assess the safety and efficacy of vaccines due to their similarity in size, genetics, reproduction (including, importantly, pregnancy and placentation), and immune repertoire and response (32, 33, 44). The larger size of the baboon compared to macaques and New World monkeys allows greater blood sampling volumes, expanding the analyses that can be performed pre- and postinfection. Also, in studies of pregnant baboons, while amniocentesis can be performed in various NHP species, the large size of the baboon allows accessing the fetal blood system via ultrasound-guided percutaneous umbilical blood sampling (PUBS) (35). Placentation is important in understanding the mechanism of vertical transfer in pregnant animals. Baboons have a monodiscoid hemochorial placenta similar to that of humans. Therefore, viral transfer and/or placental and umbilical blood flow responses to ZIKV infection may be more similar in the monodiscoid baboon placenta than in the bidiscoid placenta of rhesus macaques (with one primary and one secondary disc). The bidiscoid placenta may alter the mechanism of vertical transfer in pregnant animals and may contribute to the 100% vertical transfer seen so far in rhesus macaques (18). Baboons are also permissive to flaviviral infection and replication, including WNV, DENV, and now ZIKV. Furthermore, wild African baboons have been found to be infected with ZIKV and show a virus-specific immune response, further supporting baboons as a good translational NHP model for ZIKV infection and pathogenesis (34, 36). Our studies are significant, as they describe the first experimental infection of baboons with ZIKV. In our baboon model, the contemporary French Polynesian ZIKV isolate (H/PF/2013) caused viral infection characterized by human clinical symptoms, such as rash and conjunctivitis; viral replication in blood, urine, saliva, and CSF; persistence in the lymph nodes well after resolution in blood and other body fluids; and development of a robust ZIKV-specific IgM and IgG response and of neutralizing antibodies against ZIKV.

Adult male and female baboons showed viremia in blood between days 2 and 5 postinfection and resolution by 6 dpi, with the peak virus titer ranging in females from 2.2 × 10⁴ to 5.3 × 10⁴ RNA copies/ml blood on days 3 and 4 postinfection (low inoculum) and in males from 7.9 × 10³ to 4.0 × 10⁵ copies/ml blood (high inoculum) by 4 dpi. The difference between peak viral loads and lengths of viremia in blood between male and female baboons could be due to differences in viral dosage (10⁶ FFU in males versus 10⁴ FFU in females) and sampling days. Similar to humans, we showed prolonged detection of ZIKV RNA in urine and saliva, up to day 27 in one male (M1), and in CSF until day 11, well after resolution in the blood. We did observe ZIKV RNA in the saliva and CSF of the female (F2) with the highest peak blood viremia, suggesting initial viral seeding might be important for viral spillover and persistence in other body fluids and the CNS after clearance from the blood. In addition to fluid compartments, we found ZIKV RNA persisting in various lymph nodes in males (and, in one male, in the epididymis) at 41 days after inoculation despite the robust immune response. It is unclear how long ZIKV can persist in the lymph nodes and if other organs may contain lingering virus. In rhesus macaques, ZIKV RNA has been noted in lymph nodes for 5 to 6 weeks postinfection (22). Further, at this time, we do not know if the ZIKV found in lymph at the termination of the study was live virus. The presence of ZIKV in the epididymis of one male suggests that other reproductive organs (in particular the testes) in baboons may harbor ZIKV during the acute phase of ZIKV infection, since the virus has been detected by in situ hybridization in the testes, prostate, and seminal vesicles of cynomolgus macaques at 28 days after infection with a Thai isolate of ZIKV (22). Different strains of ZIKV may also have differences in tropism for various tissues, including reproductive organs.

The cytokine and chemokine response has been reported in ZIKV infections in patients (31 to 62 years of age) from Southeast Asia, Polynesia, and Brazil during the acute phase (less than 10 days from symptom onset) or recovery phase (22 to 62 days from symptom onset) (45). This study included both adult males and females but the influence of gender on cytokine variability was not discussed, since the sampling was unequally distributed (two females and one male for the acute phase and two males and one female for the recovery phase). During the acute phase of ZIKV infection, the authors noted significant plasma elevation of IL-1β, IL-2, IL-4, IL-6, IL-9, IL-10, IL-13, IP-10, RANTES, MIP-1α, and vascular endothelial growth factor (VEGF), while in the recovery phase, IL-1β, IL-6, IL-8, IL-10, IL-13, IP-10, RANTES, MIP-1α, MIP-1β, VEGF, fibroblast growth factor (FGF), and granulocyte-macrophage colony-stimulating factor (GM-CSF) were elevated. In the present study, we observed in the low-inoculum (female) cohort an acute-phase increase for MCP-1, IL-8, IL-2, IL-10, IL-15, and G-CSF and a trend toward increased IL-12 and IFN-γ. Several of the cytokine/chemokines measured in the human study were either not available on the NHP multiplex ELISA or the ELISA did not measure the baboon protein (IL-4, IL-9, IP-10, IL-13, RANTES, FGF, MIP-1a, MIP-1b, and GM-CSF). In the convalescent phase, we observed increases in MCP-1, IL-8, and IL-2, with a trend toward increased IL-6 and TGF-α. Of interest, IL-2 and IL-15 serve similar functions as T cell growth factors and in the conversion of naive T cells to Th1 and, as such, stimulate the early T cell response to ZIKV. IL-4 and IL-13 are involved in mediating Th2 responses, but unfortunately, the assay used was not capable of detecting these cytokines in baboon plasma. In the male baboons, we noted increases in MCP-1 and IL-8, with a trend toward increased IL-15, with decreased SCD40L in the acute phase, while in the recovery phase, we noted increases in IL-8, G-CSF, TGF-α, and SCD40L. As such, despite the higher inoculum, the acute-phase cytokine response was quite attenuated in the male baboons. One caveat to the measurement of cytokines in the present study is that the samples were taken under ketamine sedation, which may reflect a stress contribution to the findings. However, cortisol (produced in response to stress) is a potent anti-inflammatory and could have actually suppressed a more robust cytokine response, something that has to be considered for the studies in macaques, as well.

Of interest, in a study in rhesus monkeys (two females and one male), the only cytokines/chemokines that exhibited an increase in response to subcutaneous ZIKV inoculation were IL-1RA, MCP-1, IP10, and CXCL11 (ITAC) (16), while another study noted select increases in macaques in tumor necrosis factor alpha (TNF-α), IL-2, IL-10, and IL-23 (22). As such, there is some similarity in response to ZIKV between baboons and rhesus macaques; however, the baboons exhibited a much broader response, resembling what has been observed in humans. Furthermore, in the human study, day 1 was considered to be the day of presentation with symptoms, so earlier increases in cytokines/chemokines in response to ZIKV infection may have been missed, particularly in the two acute-phase human females.

Although not entirely uniform, all the baboons inoculated with ZIKV developed ZIKV-specific IgM and IgG responses. The male baboon with the highest peak viremia showed an antibody response that was not only more robust than those of the other two male baboons, but was also detected earlier. It is possible that the rapid innate and adaptive immune responses in this animal, although not as robust as those of M1, were able to limit viral replication in the blood. Lack of detectable viremia is also seen in humans. This, in combination with asymptomatic infection, which is a common occurrence with ZIKV, makes diagnosis all the more difficult, even though ZIKV may be present in other body fluids and tissues. Studies have shown that despite rapid innate and adaptive immunity to ZIKV infection, viral shedding and persistence occur in certain areas of the body, such as the CNS, lymph nodes, reproductive tissues (male and female), and body fluids (CSF and saliva), serving as viral reservoirs for future infection, disease, and sexual transmission and posing substantial risk for pregnancy. The female baboon that demonstrated the highest peak blood viremia had the most robust antibody response compared to the other two female baboons, parallel to what was observed in the male baboons. West Nile virus, with high sequence similarity to ZIKV, is endemic in the United States, including Oklahoma, and our baboon colony was maintained over the years in a facility that included outdoor housing. As such, just like the human population in the United States, our colony has animals that were exposed to WNV, likely via the mosquito vector. We know that baboons are permissive to WNV infection (34) and that baboons are naturally infected with WNV, as evidenced by preexisting antibody titers. In light of this, our one female baboon exhibiting the highest ZIKV viremia also had preexisting WNV antibody titers, while the other two female baboons did not. This female also had the highest levels of MCP-1, TNF-α, IL-2, IL-10, IL-12, IL-15, IL-1RA, and TGF-α of the three females and was the only female baboon to demonstrate spillover of virus from the blood into other body fluid compartments, such as saliva, urine, and CSF. Previous studies have demonstrated the enhancement of ZIKV disease in the presence of preexisting WNV immunity both in vitro and in vivo (46). It is possible that the increase in WNV IgG in two animals (F2 and M3) in response to ZIKV was a result of cross-reactivity of anti-WNV IgG with the ZIKV IgG ELISA; however, we did not observe cross-reactivity in these two animals on the ZIKV ELISA on day 0, and conversely, the animals seropositive for ZIKV (F1, F3, M1, and M2) were below the limit of detection on the WNV ELISA; therefore, the differences are most likely due to activation of WNV⁺ memory cells by ZIKV infection. With the known phenomenon of antibody-dependent enhancement of disease (ADE) associated with flavivirus infection, these data suggest a possible role for previous infection with WNV in enhanced ZIKV disease. However, with such a small cohort of WNV⁺ and WNV⁻ subjects, further investigation is necessary to elucidate the role that preexisting WNV immunity may play in the ZIKV baboon model. While there was no evidence for enhancement of ZIKV viremia by prior DENV infection (DENV type 1 [DENV-1] and -2) in a recent macaque study (47), prior infection of macaques with ZIKV did significantly enhance peak viremia and cytokine levels in response to DENV-2 (48), suggesting that ADE can occur between ZIKV and DENV. However, our one male baboon (M3) that was seropositive for WNV prior to ZIKV infection (and responded to ZIKV with an enhanced WNV IgG titer) had a weak IgM/IgG response to ZIKV and, in general, a very weak cytokine response to the virus. Furthermore, preexisting immunity to WNV did not seem to alter the ability of baboons to develop neutralizing antibodies compared to WNV-naive baboons in the same cohort, as evidenced by similar PRNT₅₀ endpoint titer values. Thus, the response to ZIKV in the context of WNV immunity in the baboon is likely complex and worthy of further study.

Our study establishes the olive baboon (Papio anubis) as an excellent translatable NHP model to study ZIKV infection. Baboons are naturally permissive to ZIKV disease and have an immune repertoire similar to that of humans. We were able to replicate ZIKV infection in humans in our baboon model in terms of the use of a relevant ZIKV strain associated with increased neurological symptoms and adverse fetal outcomes, pathologically relevant dosages of ZIKV, infectivity, chronicity, and immune response. Due to the similarities in placentation, genetics, immune response, and permissiveness to flaviviral infection between baboons and humans, baboons can also serve as relevant translational models for vertical and sexual transmission of ZIKV. We hope our model will play an important role in understanding the pathogenesis of this rapidly spreading and highly devastating disease and in the development of effective vaccines and therapeutics.

Go to:

MATERIALS AND METHODS

Ethics statement.

All experiments utilizing baboons were performed in compliance with guidelines established by the U.S. Animal Welfare Act for housing and care of laboratory animals, as well as the U.S. National Institutes of Health Office of Laboratory Animal Welfare Public Health Service Policy on the Humane Care and Use of Laboratory Animals in Assessment and Accreditation of Laboratory Animal Care (AAALAC) International and National accredited laboratories. All experiments were approved by the University of Oklahoma Health Sciences Center (OUHSC) Institutional Animal Care and Use Committee under animal use protocols 16-028-I and 16-140-I. All studies with ZIKV infection were performed in animal biosafety level 2 (ABSL2) containment facilities in accordance with the Centers for Disease Control and Prevention Biosafety in Microbiological Laboratories guidelines at the OUHSC. Appropriate measures were utilized to reduce potential distress, pain, and discomfort. Ketamine (10 mg/kg of body weight) was used to sedate the baboons during all procedures, which included routine blood draws performed by trained personnel. The baboons were fed standard monkey chow twice daily, as well as receiving daily food supplements (fruits). All the animals received enrichment. ZIKV-infected animals were caged separately but within visual and auditory contact with other baboons to promote social behavior and alleviate stress. At the designated times postinoculation, the animals were euthanized according to the recommendations of the American Veterinary Medical Association (2013 panel on euthanasia).

Animals.

Adult baboons (n = 3 males, aged 5 to 9 years, 25 to 30 kg body weight; n = 3 nonpregnant females, aged 5 to 8 years, 13 to 16 kg body weight) were utilized for this study.

Virus stocks and infection.

Animals were anesthetized with an intramuscular dose of ketamine before all procedures (viral inoculation and blood, swab, and body fluid collection). Nonpregnant female baboons (n = 3) and male baboons (n = 3) were infected subcutaneously in the midscapular area with a single pathologically relevant dose (volume) of 10⁴ (females) and 10⁶ (males) FFU of ZIKV derived from a French Polynesian viral isolate (H/PF/2013). The dosage used to infect the animals in our study was based on previous work done in mosquitoes carrying WNV and DENV, where it was estimated that mosquitoes carry 1 × 10⁴ to 1 × 10⁶ PFU of the virus (49), and on a study evaluating the Brazilian ZIKV range in a bite from an A. aegypti mosquito (50). Males were terminated at 41 days postinfection, and females were terminated at 43 days postinfection.

Sample collection and processing.

Animals were anesthetized before all sample collection. Sample collection days are shown in the study design and timeline (Fig. 1). Whole blood was collected into EDTA tubes. Urine was collected by direct bladder cystocentesis. Saliva was collected with a cotton roll salivette, and CSF was collected by lumbar puncture. Tissues were collected, rapidly frozen on dry ice, and stored at −80°C before processing for RNA extraction.

One-step qRT-PCR.

RNA from whole blood, saliva, urine, CSF, lymph nodes (axial, inguinal, mesentery, and submandibular; from males only), bladder, and reproductive tissues (vaginal epithelium, uterus, and ovary from nonpregnant females and left and right testes, left and right epididymis, seminal vesicle, and prostate from males) were isolated using a QIAamp cador pathogen minikit (Qiagen, Valencia, CA). ZIKV RNA was quantitated by one-step quantitative real-time reverse transcription-PCR using a QuantiTect probe RT-PCR kit (Qiagen) on an iCycler instrument (Bio-Rad). The primers and probes used for qRT-PCR were designed by Lanciotti et al. (51) (Table 2). The primers and probes were used at a concentration of 0.4 μM and 0.2 μM, respectively, and the cycling conditions used were 50°C for 30 min and 95°C for 15 min, followed by 40 cycles of 94°C for 15 s and 60°C for 1 min. The concentration of the viral RNA (copies per milliliter) was determined by interpolation on a standard curve of six 10-fold serial dilutions (10⁶ to 10¹ copies/ml) of a synthetic ZIKV RNA fragment available commercially from the ATCC (VR-3252SD). The cutoff for the limit of detection of ZIKV RNA was 1 × 10².

TABLE 2

Primer-probe sets for detection of ZIKV by one-step qRT-PCR

Set	Primers and probea	Genome position	Sequence (5′–3′)
1	ZIKV 835 Forward	835–857	TTGGTCATGATACTGCTGATTGC
	ZIKV 911 Reverse	890–911	CCTTCCACAAAGTCCCTATTGC
	ZIKV 860-FAM Probe	860–886	CGGCATACAGCATCAGGTGCATAGGAG
2	ZIKV 1086 Forward	1086–1102	CCGCTGCCCAACACAAG
	ZIKV 1162 Reverse	1162–1139	CCACTAACGTTCTTTTGCAGACAT
	ZIKV 1107-FAM Probe	1107–1137	AGCCTACCTTGACAAGCAGTCAGACACTCAA

^aFAM, 6-carboxyfluorescein.

ZIKV ELISA.

ZIKV-specific IgM and IgG antibody responses were assessed in the serum samples from males and females using commercially available anti-ZIKV IgM (ab213327; Abcam, Cambridge, MA) and IgG (Sp856C; XpressBio, Fredrick, MD) ELISA kits. Briefly, 1:100 (IgM) and 1:50 (IgG) serum dilutions were performed in duplicate and added to the precoated plates available in the kits. Male sera obtained preinfection (day 0) and 0, 4, 7, 11, 13, 27, and 34 dpi and female sera from day 0 and 4, 8, 10, 14, 21, 30, and 43 dpi were used to detect the ZIKV-specific IgM response. For the ZIKV-specific IgG response, male sera from days 0, 7, 13, 27, and 41 and female sera from days 0, 8, 14, 30, and 43 postinfection were tested. The assays were performed according to the manufacturer's instructions, and the assay was read at 450 nm for IgM and 405 nm for IgG antibodies in the serum.

WNV ELISA.

WNV-specific IgG antibody responses were assessed in the serum samples in males and nonpregnant females using a simian West Nile ELISA kit (SP807C; XpressBio, Fredrick, MD). Briefly, a 1:50 serum dilution was performed in duplicate and added to the precoated plates in the kit. Sera collected preinfection (day 0) and days 7, 13, 27, and 41 postinfection from males and day 0 and days 8, 14, 30, and 43 postinfection from females were tested for WNV IgG response. The assays were performed according to the manufacturer's instructions, and the assay was read at 405 nm.

Plaque reduction neutralization test.

A PRNT was used to assess serum samples for ZIKV neutralizing antibodies (52). Vero cells (ATCC CCL-81) were maintained in Dulbecco's modified Eagle's medium (DMEM) (supplemented with 10% heat-inactivated fetal bovine serum [FBS], 1× antibiotic/antimycotic), seeded in 12-well plates (2 × 10⁵ cells/well; 37°C; 5% CO₂), and incubated at for approximately 24 h until 80 to 100% confluent. Baboon sera were heat inactivated (56°C; 30 min) and then serially diluted 2-fold in medium, followed by addition of 200 PFU of the French Polynesian ZIKV isolate (H/PF/2013). Samples were vortexed and incubated in a 37°C water bath for 1 h. The medium was then removed from each well and replaced with the virus-serum mixture, followed by incubation at 37°C for 1 h with intermittent rocking of the plates every 20 min. Control wells included inoculation with 200 PFU of ZIKV with no serum added to determine total plaques, as well as control wells without virus. A 1% carboxymethyl cellulose overlay was then added without removing the inoculum, and the plates were incubated at 37°C for approximately 72 h. The cells were then fixed with 1% paraformaldehyde for 1 h, after which the overlay was removed and the cells were stained with 1% crystal violet. The PRNT₅₀ was calculated by visually counting plaques. All samples were tested in duplicate.

Serum cytokine analysis.

A nonhuman primate cytokine magnetic bead 23-plex panel (catalog number PRCYTOMG-40k; EMD Millipore, Billerica, MA) was used to quantify cytokines and chemokines in sera from male and female baboons. The assays were performed according to the manufacturer's instructions. Briefly, 25 μl of serum samples was incubated with 23-plex premixed antibody-conjugated magnetic polystyrene beads overnight in a 96-well plate. The beads were washed in wash buffer and incubated in biotinylated detection antibody for 1 h, followed by streptavidin-phycoerythrin incubation for 30 min. After the final wash, cytokines were detected and analyzed with a BioPlex-200 detection system (Bio-Rad). High- and low-concentration control sera (supplied in the kits) were run with each ELISA; the interassay coefficient of variation was ≤10%. A paired t test was used to test for differences between preinfection (day 0) and the peak cytokine obtained in the acute phase (≤10 dpi) and latent phase (>14 dpi) for each baboon, since individual starting concentrations were highly variable between animals. Prism (GraphPad, Inc.; v7) was used for statistical analysis. Due to the limited animal numbers per group (n = 3), variability in the day postinfection of the peak response for each cytokine precluded the use of repeated-measures analysis of variance (ANOVA).

IFN-γ ELISPOT assay.

PBMCs were stimulated with recombinant protein from Zika envelope (1 μg/ml) or NS1 (1 μg/ml) or PHA (1 μg/ml) as a positive control to determine the numbers of IFN-γ-producing cells by ELISPOT assay using the methodology reported previously (53,–56). Briefly, aliquots of PBMCs (10⁵/well) were seeded in duplicate wells of 96-well plates (polyvinylidene difluoride-backed plates [MAIP S 45; Millipore, Bedford, MA]) precoated with the primary IFN-γ antibody, and the lymphocytes were stimulated with the different mitogens. After incubation for 30 to 36 h at 37°C, the cells were removed and the wells were thoroughly washed with PBS and developed according to the protocol provided by the manufacturer. The results are expressed as IFN-γ spot-forming cells (SFCs) per 10⁵ PBMCs after subtraction of the duplicate wells with medium only (negative control) and were considered positive if greater than twice the background and if there were more than 5 SFCs/10⁵ PBMCs. Both recombinant proteins from Zika envelope and NS1 were purchased from My BioSource (San Diego, CA).

Go to:

ACKNOWLEDGMENTS

We thank the Oklahoma Baboon Research Resource for providing baboons; James J. Tomasek, Vice President for Research, University of Oklahoma Health Sciences Center, for support of the project; and Helen Lazear, University of North Carolina Chapel Hill, for providing ZIKV stocks.

This study was supported by NIH R21 AI129560, NIH R21 NS103772, NIH P40 OD010988, and funds from the University of Oklahoma Health Sciences Center Vice President for Research.

J.F.P., D.A.M., and S.G. designed the experiments, analyzed the data, and drafted the manuscript. S.G. developed and performed viral load assays, ZIKV IgM ELISAs, and cytokine assays. A.N.P., J.F.P, R.F.W. and J.P.D. performed the baboon necropsies and helped with sample collection. N.R. coordinated baboon tissue sampling for processing and performed blood chemistries, CBC panel, and ZIKV and WNV IgG ELISAs. H.N. and K.H. performed the PRNTs. B.N. and P.N. performed the IFN-γ ELISPOT assays, and D.P.D. assisted with viral load assays, data analysis, and drafting the manuscript.

We declare that we have no competing financial interests.

Go to:

REFERENCES

1. Dick GW. 1952. Zika virus. II. Pathogenicity and physical properties. Trans R Soc Trop Med Hyg46:521–534. doi:10.1016/0035-9203(52)90043-6. [PubMed] [CrossRef] [Google Scholar]
2. Dick GW, Kitchen SF, Haddow AJ. 1952. Zika virus. I. Isolations and serological specificity. Trans R Soc Trop Med Hyg46:509–520. doi:10.1016/0035-9203(52)90042-4. [PubMed] [CrossRef] [Google Scholar]
3. Marchette NJ, Garcia R, Rudnick A. 1969. Isolation of Zika virus from Aedes aegypti mosquitoes in Malaysia. Am J Trop Med Hyg18:411–415. doi:10.4269/ajtmh.1969.18.411. [PubMed] [CrossRef] [Google Scholar]
4. Brasil P, Pereira JP Jr, Moreira ME, Ribeiro Nogueira RM, Damasceno L, Wakimoto M, Rabello RS, Valderramos SG, Halai UA, Salles TS, Zin AA, Horovitz D, Daltro P, Boechat M, Raja Gabaglia C, Carvalho de Sequeira P, Pilotto JH, Medialdea-Carrera R, Cotrim da Cunha D, Abreu de Carvalho LM, Pone M, Machado Siqueira A, Calvet GA, Rodrigues Baiao AE, Neves ES, Nassar de Carvalho PR, Hasue RH, Marschik PB, Einspieler C, Janzen C, Cherry JD, Bispo de Filippis AM, Nielsen-Saines K. 2016. Zika virus infection in pregnant women in Rio de Janeiro. N Engl J Med375:2321–2334. doi:10.1056/NEJMoa1602412. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
5. D'Ortenzio E, Matheron S, Yazdanpanah Y, de Lamballerie X, Hubert B, Piorkowski G, Maquart M, Descamps D, Damond F, Leparc-Goffart I. 2016. Evidence of sexual transmission of Zika virus. N Engl J Med374:2195–2198. doi:10.1056/NEJMc1604449. [PubMed] [CrossRef] [Google Scholar]
6. Turmel JM, Abgueguen P, Hubert B, Vandamme YM, Maquart M, Le Guillou-Guillemette H, Leparc-Goffart I. 2016. Late sexual transmission of Zika virus related to persistence in the semen. Lancet387:2501. doi:10.1016/S0140-6736(16)30775-9. [PubMed] [CrossRef] [Google Scholar]
7. Oehler E, Watrin L, Larre P, Leparc-Goffart I, Lastere S, Valour F, Baudouin L, Mallet H, Musso D, Ghawche F. 2014. Zika virus infection complicated by Guillain-Barre syndrome—case report, French Polynesia, December 2013. Euro Surveill19:20720. doi:10.2807/1560-7917.ES2014.19.9.20720. [PubMed] [CrossRef] [Google Scholar]
8. Duffy MR, Chen TH, Hancock WT, Powers AM, Kool JL, Lanciotti RS, Pretrick M, Marfel M, Holzbauer S, Dubray C, Guillaumot L, Griggs A, Bel M, Lambert AJ, Laven J, Kosoy O, Panella A, Biggerstaff BJ, Fischer M, Hayes EB. 2009. Zika virus outbreak on Yap Island, Federated States of Micronesia. N Engl J Med360:2536–2543. doi:10.1056/NEJMoa0805715. [PubMed] [CrossRef] [Google Scholar]
9. European Centre for Disease Prevention and Control. 2016. Rapid risk assessment. Zika virus disease epidemic: Sixth update, 20 May 2016 ECDC, Stockholm, Sweden. [Google Scholar]
10. Cugola FR, Fernandes IR, Russo FB, Freitas BC, Dias JL, Guimaraes KP, Benazzato C, Almeida N, Pignatari GC, Romero S, Polonio CM, Cunha I, Freitas CL, Brandao WN, Rossato C, Andrade DG, Faria Dde P, Garcez AT, Buchpigel CA, Braconi CT, Mendes E, Sall AA, Zanotto PM, Peron JP, Muotri AR, Beltrao-Braga PC. 2016. The Brazilian Zika virus strain causes birth defects in experimental models. Nature534:267–271. doi:10.1038/nature18296. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
11. Lazear HM, Govero J, Smith AM, Platt DJ, Fernandez E, Miner JJ, Diamond MS. 2016. A mouse model of Zika virus pathogenesis. Cell Host Microbe19:720–730. doi:10.1016/j.chom.2016.03.010. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
12. Miner JJ, Cao B, Govero J, Smith AM, Fernandez E, Cabrera OH, Garber C, Noll M, Klein RS, Noguchi KK, Mysorekar IU, Diamond MS. 2016. Zika virus infection during pregnancy in mice causes placental damage and fetal demise. Cell165:1081–1091. doi:10.1016/j.cell.2016.05.008. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
13. Adibi JJ, Marques ETA, Cartus A, Beigi RH. 2016. Teratogenic effects of the Zika virus and the role of the placenta. Lancet387:1587–1590. doi:10.1016/S0140-6736(16)00650-4. [PubMed] [CrossRef] [Google Scholar]
14. França GVA, Schuler-Faccini L, Oliveira WK, Henriques CMP, Carmo EH, Pedi VD, Nunes ML, Castro MC, Serruya S, Silveira MF, Barros FC, Victora CG. 2016. Congenital Zika virus syndrome in Brazil: a case series of the first 1501 live births with complete investigation. Lancet388:891–897. doi:10.1016/S0140-6736(16)30902-3. [PubMed] [CrossRef] [Google Scholar]
15. Dudley DM, Aliota MT, Mohr EL, Weiler AM, Lehrer-Brey G, Weisgrau KL, Mohns MS, Breitbach ME, Rasheed MN, Newman CM, Gellerup DD, Moncla LH, Post J, Schultz-Darken N, Schotzko ML, Hayes JM, Eudailey JA, Moody MA, Permar SR, O'Connor SL, Rakasz EG, Simmons HA, Capuano S, Golos TG, Osorio JE, Friedrich TC, O'Connor DH. 2016. A rhesus macaque model of Asian-lineage Zika virus infection. Nat Commun7:12204. doi:10.1038/ncomms12204. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
16. Hirsch AJ, Smith JL, Haese NN, Broeckel RM, Parkins CJ, Kreklywich C, DeFilippis VR, Denton M, Smith PP, Messer WB, Colgin LM, Ducore RM, Grigsby PL, Hennebold JD, Swanson T, Legasse AW, Axthelm MK, MacAllister R, Wiley CA, Nelson JA, Streblow DN. 2017. Zika virus infection of rhesus macaques leads to viral persistence in multiple tissues. PLoS Pathog13:e1006219. doi:10.1371/journal.ppat.1006219. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
17. Osuna CE, Lim SY, Deleage C, Griffin BD, Stein D, Schroeder LT, Omange R, Best K, Luo M, Hraber PT, Andersen-Elyard H, Ojeda EF, Huang S, Vanlandingham DL, Higgs S, Perelson AS, Estes JD, Safronetz D, Lewis MG, Whitney JB. 2016. Zika viral dynamics and shedding in rhesus and cynomolgus macaques. Nat Med22:1448–1455. doi:10.1038/nm.4206. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
18. Nguyen SM, Antony KM, Dudley DM, Kohn S, Simmons HA, Wolfe B, Salamat MS, Teixeira LBC, Wiepz GJ, Thoong TH, Aliota MT, Weiler AM, Barry GL, Weisgrau KL, Vosler LJ, Mohns MS, Breitbach ME, Stewart LM, Rasheed MN, Newman CM, Graham ME, Wieben OE, Turski PA, Johnson KM, Post J, Hayes JM, Schultz-Darken N, Schotzko ML, Eudailey JA, Permar SR, Rakasz EG, Mohr EL, Capuano S, Tarantal AF, Osorio JE, O'Connor SL, Friedrich TC, O'Connor DH, Golos TG. 2017. Highly efficient maternal-fetal Zika virus transmission in pregnant rhesus macaques. PLoS Pathog13:e1006378. doi:10.1371/journal.ppat.1006378. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
19. Haddow AD, Nalca A, Rossi FD, Miller LJ, Wiley MR, Perez-Sautu U, Washington SC, Norris SL, Wollen-Roberts SE, Shamblin JD, Kimmel AE, Bloomfield HA, Valdez SM, Sprague TR, Principe LM, Bellanca SA, Cinkovich SS, Lugo-Roman L, Cazares LH, Pratt WD, Palacios GF, Bavari S, Pitt ML, Nasar F. 2017. High infection rates for adult macaques after intravaginal or intrarectal inoculation with Zika virus. Emerg Infect Dis23:1274–1281. doi:10.3201/eid2308.170036. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
20. Newman CM, Dudley DM, Aliota MT, Weiler AM, Barry GL, Mohns MS, Breitbach ME, Stewart LM, Buechler CR, Graham ME, Post J, Schultz-Darken N, Peterson E, Newton W, Mohr EL, Capuano S III, O'Connor DH, Friedrich TC. 2017. Oropharyngeal mucosal transmission of Zika virus in rhesus macaques. Nat Commun8:169. doi:10.1038/s41467-017-00246-8. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
21. Carroll T, Lo M, Lanteri M, Dutra J, Zarbock K, Silveira P, Rourke T, Ma ZM, Fritts L, O'Connor S, Busch M, Miller CJ. 2017. Zika virus preferentially replicates in the female reproductive tract after vaginal inoculation of rhesus macaques. PLoS Pathog13:e1006537. doi:10.1371/journal.ppat.1006537. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
22. Aid M, Abbink P, Larocca RA, Boyd M, Nityanandam R, Nanayakkara O, Martinot AJ, Moseley ET, Blass E, Borducchi EN, Chandrashekar A, Brinkman AL, Molloy K, Jetton D, Tartaglia LJ, Liu J, Best K, Perelson AS, De La Barrera RA, Lewis MG, Barouch DH. 2017. Zika virus persistence in the central nervous system and lymph nodes of rhesus monkeys. Cell169:610–620.e14. doi:10.1016/j.cell.2017.04.008. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
23. Silveira ELV, Rogers KA, Gumber S, Amancha P, Xiao P, Woollard SM, Byrareddy SN, Teixeira MM, Villinger F. 2017. Immune cell dynamics in rhesus macaques infected with a Brazilian strain of Zika virus. J Immunol199:1003–1011. doi:10.4049/jimmunol.1700256. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
24. Coffey LL, Pesavento PA, Keesler RI, Singapuri A, Watanabe J, Watanabe R, Yee J, Bliss-Moreau E, Cruzen C, Christe KL, Reader JR, von Morgenland W, Gibbons AM, Allen AM, Linnen J, Gao K, Delwart E, Simmons G, Stone M, Lanteri M, Bakkour S, Busch M, Morrison J, Van Rompay KK. 2017. Zika virus tissue and blood compartmentalization in acute infection of rhesus macaques. PLoS One12:e0171148. doi:10.1371/journal.pone.0171148. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
25. Koide F, Goebel S, Snyder B, Walters KB, Gast A, Hagelin K, Kalkeri R, Rayner J. 2016. Development of a Zika virus infection model in cynomolgus macaques. Front Microbiol7:2028. doi:10.3389/fmicb.2016.02028. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
26. Adams Waldorf KM, Stencel-Baerenwald JE, Kapur RP, Studholme C, Boldenow E, Vornhagen J, Baldessari A, Dighe MK, Thiel J, Merillat S, Armistead B, Tisoncik-Go J, Green RR, Davis MA, Dewey EC, Fairgrieve MR, Gatenby JC, Richards T, Garden GA, Diamond MS, Juul SE, Grant RF, Kuller L, Shaw DW, Ogle J, Gough GM, Lee W, English C, Hevner RF, Dobyns WB, Gale M Jr, Rajagopal L. 2016. Fetal brain lesions after subcutaneous inoculation of Zika virus in a pregnant nonhuman primate. Nat Med22:1256–1259. doi:10.1038/nm.4193. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
27. Mohr EL, Block LN, Newman CM, Stewart LM, Koenig M, Semler M, Breitbach ME, Teixeira LBC, Zeng X, Weiler AM, Barry GL, Thoong TH, Wiepz GJ, Dudley DM, Simmons HA, Mejia A, Morgan TK, Salamat MS, Kohn S, Antony KM, Aliota MT, Mohns MS, Hayes JM, Schultz-Darken N, Schotzko ML, Peterson E, Capuano S III, Osorio JE, O'Connor SL, Friedrich TC, O'Connor DH, Golos TG. 2018. Ocular and uteroplacental pathology in a macaque pregnancy with congenital Zika virus infection. PLoS One13:e0190617. doi:10.1371/journal.pone.0190617. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
28. Hirsch AJ, Roberts VHJ, Grigsby PL, Haese N, Schabel MC, Wang X, Lo JO, Liu Z, Kroenke CD, Smith JL, Kelleher M, Broeckel R, Kreklywich CN, Parkins CJ, Denton M, Smith P, DeFilippis V, Messer W, Nelson JA, Hennebold JD, Grafe M, Colgin L, Lewis A, Ducore R, Swanson T, Legasse AW, Axthelm MK, MacAllister R, Moses AV, Morgan TK, Frias AE, Streblow DN. 2018. Zika virus infection in pregnant rhesus macaques causes placental dysfunction and immunopathology. Nat Commun9:263. doi:10.1038/s41467-017-02499-9. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
29. Magnani DM, Rogers TF, Maness NJ, Grubaugh ND, Beutler N, Bailey VK, Gonzalez-Nieto L, Gutman MJ, Pedreno-Lopez N, Kwal JM, Ricciardi MJ, Myers TA, Julander JG, Bohm RP, Gilbert MH, Schiro F, Aye PP, Blair RV, Martins MA, Falkenstein KP, Kaur A, Curry CL, Kallas EG, Desrosiers RC, Goldschmidt-Clermont PJ, Whitehead SS, Andersen KG, Bonaldo MC, Lackner AA, Panganiban AT, Burton DR, Watkins DI. 2018. Fetal demise and failed antibody therapy during Zika virus infection of pregnant macaques. Nat Commun9:1624. doi:10.1038/s41467-018-04056-4. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
30. Chiu CY, Sanchez-San Martin C, Bouquet J, Li T, Yagi S, Tamhankar M, Hodara VL, Parodi LM, Somasekar S, Yu G, Giavedoni LD, Tardif S, Patterson J. 2017. Experimental Zika virus inoculation in a New World monkey model reproduces key features of the human infection. Sci Rep7:17126. doi:10.1038/s41598-017-17067-w. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
31. Vanchiere JA, Ruiz JC, Brady AG, Kuehl TJ, Williams LE, Baze WB, Wilkerson GK, Nehete PN, McClure GB, Rogers DL, Rossi SL, Azar SR, Roundy CM, Weaver SC, Vasilakis N, Simmons JH, Abee CR. 2018. Experimental Zika virus infection of neotropical primates. Am J Trop Med Hyg98:173–177. doi:10.4269/ajtmh.17-0322. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
32. Shearer MH, Dark RD, Chodosh J, Kennedy RC. 1999. Comparison and characterization of immunoglobulin G subclasses among primate species. Clin Diagn Lab Immunol6:953–958. [PMC free article] [PubMed] [Google Scholar]
33. Shearer MH, Lucas AH, Anderson PW, Carey KD, Jenson HB, Chanh TC, Stanley JR, Kennedy RC. 1997. The baboon as a nonhuman primate model for assessing the effects of maternal immunization with Haemophilus influenzae type b polysaccharide vaccines. Infect Immun65:3267–3270. [PMC free article] [PubMed] [Google Scholar]
34. Wolf RF, Papin JF, Hines-Boykin R, Chavez-Suarez M, White GL, Sakalian M, Dittmer DP. 2006. Baboon model for West Nile Virus infection and vaccine evaluation. Virology355:44–51. doi:10.1016/j.virol.2006.06.033. [PubMed] [CrossRef] [Google Scholar]
35. Watts AM, Stanley JR, Shearer MH, Hefty PS, Kennedy RC. 1999. Fetal immunization of baboons induces a fetal-specific antibody response. Nat Med5:427–430. doi:10.1038/7426. [PubMed] [CrossRef] [Google Scholar]
36. Buechler CR, Bailey AL, Weiler AM, Barry GL, Breitbach ME, Stewart LM, Jasinska AJ, Freimer NB, Apetrei C, Phillips-Conroy JE, Jolly CJ, Rogers J, Friedrich TC, O'Connor DH. 2017. Seroprevalence of Zika virus in wild African green monkeys and baboons. mSphere2:e00392-. doi:10.1128/mSphere.00392-16. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
37. Ye Q, Liu ZY, Han JF, Jiang T, Li XF, Qin CF. 2016. Genomic characterization and phylogenetic analysis of Zika virus circulating in the Americas. Infect Genet Evol43:43–49. doi:10.1016/j.meegid.2016.05.004. [PubMed] [CrossRef] [Google Scholar]
38. Enfissi A, Codrington J, Roosblad J, Kazanji M, Rousset D. 2016. Zika virus genome from the Americas. Lancet387:227–228. doi:10.1016/S0140-6736(16)00003-9. [PubMed] [CrossRef] [Google Scholar]
39. Wang L, Valderramos SG, Wu A, Ouyang S, Li C, Brasil P, Bonaldo M, Coates T, Nielsen-Saines K, Jiang T, Aliyari R, Cheng G. 2016. From mosquitos to humans: genetic evolution of Zika virus. Cell Host Microbe19:561–565. doi:10.1016/j.chom.2016.04.006. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
40. Yuan L, Huang XY, Liu ZY, Zhang F, Zhu XL, Yu JY, Ji X, Xu YP, Li G, Li C, Wang HJ, Deng YQ, Wu M, Cheng ML, Ye Q, Xie DY, Li XF, Wang X, Shi W, Hu B, Shi PY, Xu Z, Qin CF. 2017. A single mutation in the prM protein of Zika virus contributes to fetal microcephaly. Science358:933–936. doi:10.1126/science.aam7120. [PubMed] [CrossRef] [Google Scholar]
41. Harewood WJ, Gillin A, Hennessy A, Armistead J, Horvath JS, Tiller DJ. 1999. Biochemistry and haematology values for the baboon (Papio hamadryas): the effects of sex, growth, development and age. J Med Primatol28:19–31. doi:10.1111/j.1600-0684.1999.tb00085.x. [PubMed] [CrossRef] [Google Scholar]
42. Morrison TE, Diamond MS. 2017. Animal models of Zika virus infection, pathogenesis, and immunity. J Virol91:e00009-. doi:10.1128/JVI.00009-17. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
43. Smith DG. 2005. Genetic characterization of Indian-origin and Chinese-origin rhesus macaques (Macaca mulatta). Comp Med55:227–230. [PubMed] [Google Scholar]
44. Damian RT, Greene ND, Kalter SS. 1971. IgG subclasses in the baboon (Papio cynocephalus). J Immunol106:246–257. [PubMed] [Google Scholar]
45. Tappe D, Perez-Giron JV, Zammarchi L, Rissland J, Ferreira DF, Jaenisch T, Gomez-Medina S, Gunther S, Bartoloni A, Munoz-Fontela C, Schmidt-Chanasit J. 2016. Cytokine kinetics of Zika virus-infected patients from acute to reconvalescent phase. Med Microbiol Immunol205:269–273. doi:10.1007/s00430-015-0445-7. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
46. Bardina SV, Bunduc P, Tripathi S, Duehr J, Frere JJ, Brown JA, Nachbagauer R, Foster GA, Krysztof D, Tortorella D, Stramer SL, Garcia-Sastre A, Krammer F, Lim JK. 2017. Enhancement of Zika virus pathogenesis by preexisting antiflavivirus immunity. Science356:175–180. doi:10.1126/science.aal4365. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
47. Pantoja P, Perez-Guzman EX, Rodriguez IV, White LJ, Gonzalez O, Serrano C, Giavedoni L, Hodara V, Cruz L, Arana T, Martinez MI, Hassert MA, Brien JD, Pinto AK, de Silva A, Sariol CA. 2017. Zika virus pathogenesis in rhesus macaques is unaffected by pre-existing immunity to dengue virus. Nat Commun8:15674. doi:10.1038/ncomms15674. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
48. George J, Valiant WG, Mattapallil MJ, Walker M, Huang YS, Vanlandingham DL, Misamore J, Greenhouse J, Weiss DE, Verthelyi D, Higgs S, Andersen H, Lewis MG, Mattapallil JJ. 2017. Prior exposure to Zika virus significantly enhances peak dengue-2 viremia in rhesus macaques. Sci Rep7:10498. doi:10.1038/s41598-017-10901-1. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
49. Styer LM, Kent KA, Albright RG, Bennett CJ, Kramer LD, Bernard KA. 2007. Mosquitoes inoculate high doses of West Nile virus as they probe and feed on live hosts. PLoS Pathog3:1262–1270. doi:10.1371/journal.ppat.0030132. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
50. Dutra HL, Rocha MN, Dias FB, Mansur SB, Caragata EP, Moreira LA. 2016. Wolbachia blocks currently circulating Zika virus isolates in Brazilian Aedes aegypti mosquitoes. Cell Host Microbe19:771–774. doi:10.1016/j.chom.2016.04.021. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
51. Lanciotti RS, Kosoy OL, Laven JJ, Velez JO, Lambert AJ, Johnson AJ, Stanfield SM, Duffy MR. 2008. Genetic and serologic properties of Zika virus associated with an epidemic, Yap State, Micronesia, 2007. Emerg Infect Dis14:1232–1239. doi:10.3201/eid1408.080287. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
52. World Health Organization. 2007. Guidelines for plaque-reduction neutralization testing of human antibodies to dengue viruses. WHO/IVB/07 07. WHO, Geneva, Switzerland. [Google Scholar]
53. Nehete P, Magden E, Nehete B, Williams L, Abee C, Sastry K. 2017. Age- and sex-associated differences in phenotypic and functional characteristics of peripheral blood lymphocytes in chimpanzees (Pan troglodytes). J Am Assoc Lab Anim Sci56:509–519. [PubMed] [Google Scholar]
54. Nehete PN, Hanley PW, Nehete BP, Yang G, Ruiz JC, Williams L, Abee CR, Sastry KJ. 2013. Phenotypic and functional characterization of lymphocytes from different age groups of Bolivian squirrel monkeys (Saimiri boliviensis boliviensis). PLoS One8:e79836. doi:10.1371/journal.pone.0079836. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
55. Nehete PN, Nehete BP, Chitta S, Williams LE, Abee CR. 2017. Phenotypic and functional characterization of peripheral blood lymphocytes from various age- and sex-specific groups of owl monkeys (Aotus nancymaae). Comp Med67:67–78. [PMC free article] [PubMed] [Google Scholar]
56. Nehete PN, Shelton KA, Nehete BP, Chitta S, Williams LE, Schapiro SJ, Abee CR. 2017. Effects of transportation, relocation, and acclimation on phenotypes and functional characteristics of peripheral blood lymphocytes in rhesus monkeys (Macaca mulatta). PLoS One12:e0188694. doi:10.1371/journal.pone.0188694. [PMC free article] [PubMed] [CrossRef] [Google Scholar]

Articles from Journal of Virology are provided here courtesy of American Society for Microbiology (ASM)