Enzootic vs Epizootic?

Enzootic vs Epizootic?

We are searching data for your request:

Forums and discussions:
Manuals and reference books:
Data from registers:
Wait the end of the search in all databases.
Upon completion, a link will appear to access the found materials.

I'm studying microbiology and I see these words - epizootic and enzootic, often but there are no clear explanations for them online. Can someone help please?

Enzootic and epizootic are analogous to endemic and epidemic, respectively.

Enzootic means something that affects a population of non-human animals in a limited spatio-temporal region whereas epizootic is relatively more widespread.

Endemic and epidemic are more general terms and do not, technically, apply to only humans.

Epizootic to enzootic transition of a fungal disease in tropical Andean frogs: Are surviving species still susceptible?

The fungal pathogen Batrachochytrium dendrobatidis (Bd), which causes the disease chytridiomycosis, has been linked to catastrophic amphibian declines throughout the world. Amphibians differ in their vulnerability to chytridiomycosis some species experience epizootics followed by collapse while others exhibit stable host/pathogen dynamics where most amphibian hosts survive in the presence of Bd (e.g., in the enzootic state). Little is known about the factors that drive the transition between the two disease states within a community, or whether populations of species that survived the initial epizootic are stable, yet this information is essential for conservation and theory. Our study focuses on a diverse Peruvian amphibian community that experienced a Bd-caused collapse. We explore host/Bd dynamics of eight surviving species a decade after the mass extinction by using population level disease metrics and Bd-susceptibility trials. We found that three of the eight species continue to be susceptible to Bd, and that their populations are declining. Only one species is growing in numbers and it was non-susceptible in our trials. Our study suggests that some species remain vulnerable to Bd and exhibit ongoing population declines in enzootic systems where Bd-host dynamics are assumed to be stable.

Conflict of interest statement

Competing Interests: The authors have declared that no competing interests exist.


Fig 1. Variation in Bd infection prevalence…

Fig 1. Variation in Bd infection prevalence for eight species of frogs in the montane…

Fig 2. Differences in survival and Bd…

Fig 2. Differences in survival and Bd infection intensity in two species of marsupial frogs…

Fig 3. The gladiator frog Hypsiboas armatus…

Fig 3. The gladiator frog Hypsiboas armatus (Hylidae) and the Cusco Andes frog Psychrophrynella usurpator…

Fig 4. Differences in survival and Bd…

Fig 4. Differences in survival and Bd infection intensity in two non-susceptible species of terrestrial-breeding…

Fig 5. Differences in survival and Bd…

Fig 5. Differences in survival and Bd infection intensity in two susceptible species of terrestrial-breeding…

Fig 6. Elevational ranges and summary of…

Fig 6. Elevational ranges and summary of findings for the species examined in this study.


Mark-Recapture Study: Survival and Persistence with Bd.

In a 5-year study, a total of 392 adult R. sierrae were tagged at three persistent Bd-infected sites. Each site had consistently small R. sierrae population counts the average number of adult frogs observed per visit was only 8 (± 1 SE) at site 1, 23 (± 4) at site 2, and 31 (± 5) at site 3. But reproduction occurred each year, and tadpoles were present every year. On average, 60% of the adult frogs at site 1, 76% of those at site 2, and 74% of those at site 3 were infected on each sample date (Fig. 1 A–C). No consistent seasonal trend in the prevalence of adult infection was observed the prevalence increased significantly from the start to the end of the summer season only at site 3 [logistic regression: logit(prevalence at site 3) = 0.18 + 0.01 · days since June 1 of the year P < 0.01].

Bd load data from R. sierrae at sites with enzootic infections. (A–C) Infection prevalence and distribution of Bd loads in adult R. sierrae at three sites over 5 years. The number of individuals swabbed is shown at the top of each bar. The distribution of Bd load, as measured by the numbers of zoospores per swab (Zswab) as estimated by real-time PCR is shown in the colored bars. (D) Comparison of the fungal load in young tadpoles (Gosner stage ≤ 37), old tadpoles (Gosner stage 38–41), metamorphs, and adults at the three sites for all dates combined. Bd loads on subadults (postmetamorphic individuals with a snout-to-vent length of 35–40 mm) was not significantly different from those on adult frogs, and subadults and adults are combined in the figure. (E) Examples of changes in Bd load through time for individually marked adult R. sierrae. Shown are Bd loads for six of the individuals at site 3 that were captured in multiple years.

Infected adult frogs frequently lost and gained Bd infection through time (Fig. 1E). Of the 215 frogs that were captured more than once, 95.0% were infected during at least one of the capture events, but 38.6% transitioned from being infected to being uninfected at least once during the study, and 41.9% made the transition from uninfected to infected. Infected adults had low infection intensities as measured by the number of zoospore equivalents detected by real-time PCR on a standardized skin swab, termed “Bd load” (Fig. 1 A–E). The mean Bd load on infected adults across the three sites was 220 (± 81) zoospore equivalents per standardized swab, with a median value of only 20.5 zoospore equivalents (including all individuals with a snout-to-vent length ≥ 40 mm). These levels are much lower than the Bd loads reported in a recent study of R. muscosa/R. sierrae populations elsewhere in the Sierra Nevada during massive die-offs (16). In that study, after the first appearance of Bd, zoospore load increased approximately exponentially until the average zoospore load reached 10 4 to 10 5 per standardized swab, at which point the frog populations collapsed, in many cases to extirpation. In the current study, only two adult frogs at any of the persistent sites were ever observed to have a Bd load exceeding 10 4 per standardized swab. In contrast, the tadpoles and recently metamorphed individuals had significantly higher Bd loads compared with adults [Fig. 1D ANOVA on log-transformed Bd loads on individuals with a Bd load >0, P < 0.01 tadpoles: mean, 4,013 ± 289, median, 1,744 zoospores per standardized swab metamorphs (individuals with Gosner stage 45 or 46 with a snout-to-vent length <35 mm): mean, 4,913 ± 820 median, 631 zoospores per standardized swab).

The most important result from the multistate mark-recapture analysis (32) was that Bd infection status had no detectable effect on adult frog survival at these persistent sites (likelihood ratio test comparing a model in which the survival probability depends on infection status and site with one in which the survival probability depends only on site χ 2 = 1.5, df = 3, P > 0.6). The survival rates were significantly different between the sites, however (likelihood ratio test comparing models in which survival probability varied between sites with model with a constant survival rate χ 2 = 15.9, df = 2, P < 0.001), with best estimates of annual adult survival probabilities of 48.2% at site 1, 79.4% at site 2, and 86.5% at site 3. The “best model” from the mark-recapture analysis was one in which the adult frog survival probability depended only on site and not on infection status or time, the state transition probabilities depended on infection status (with the monthly transition rate from uninfected to infected higher at each site than the monthly transition rate from infected to uninfected) and site, and the capture probabilities depended on site and time (due to variability in survey conditions and capture effort). The model comparisons, best estimates, and 95% confidence intervals (CIs) for all model parameters are provided in SI Methods.

Bd Load Model.

The Bd load model follows the number of zoospores in a zoospore pool (i.e., a lake or pond containing a population of frogs), and the number of sporangia on each frog (Fig. 2A). The model describing the zoospore dynamics in a single frog in a zoospore pool is a set of linear ordinary differential equations, and thus the only two potential outcomes are exponential growth (resulting in death of the frog when the number of sporangia reaches the maximum level that a frog can tolerate, Smax) or exponential decline (loss of infection) at rate λ (the dominant eigenvalue of the linear system). Exponential growth of sporangia on a single frog (λ > 0) occurs if the zoospore release and reinfection rates are greater than the loss rates of sporangia from the frog skin. Local environmental conditions, including temperature, humidity, and/or water flow rate are likely to affect the reinfection and loss rates, such that individuals of a given species of amphibian may die due to the infection in some environmental conditions, but lose the infection in other conditions (Fig. 2B).

The Bd load model, and results from the deterministic version within a year. (A) Diagram of the within-host/zoospore pool model. (B) Growth rate of Bd on individual frogs (λ), as a function of f, the fraction of zoospores that reencounter the host from which they were released, and ν, the fraction of zoospores that successfully infect the frog skin on encountering a host. Other parameters are V = 1 unit volume, γ = 0.01 unit volume · day −1 , μ = 1 day −1 , η = 17.5 zoospores · day −1 , and σ = 0.2 day −1 . (C) Growth rate, λ, of Bd in the zoospore pool and time to reach Smax = 10,000 sporangia as a function of frog density. Parameters are as in B, with ν = 0.05 and f = 0.05. With few frogs present, the growth of Bd on each frog is negative, and all frogs will clear the infection. Above a density of

20 frogs, the Bd growth rate is positive on each frog. Also shown is the number of days that it takes for the density of sporangia on each frog to reach Smax.

The growth rate of Bd (λ) is an increasing function of frog density (Fig. 2C). For encounter and reinfection rates for which Bd on a single frog has a negative growth rate, there is a threshold frog population size, NT, above which the Bd growth rate shifts from negative to positive. Increasing the number of frogs further above NT also decreases the time needed for the density of sporangia on each frog to reach Smax (i.e., the time to death due to chytridiomycosis decreases). Thus, the inclusion of Bd load dynamics in this model can explain the rapid mortality due to Bd in some frog populations (especially when Bd first invades large, uninfected populations), but survival and potential recovery of frogs in others. The death of frogs when their Bd load exceeds Smax creates a negative feedback, such that it is possible for disease-induced mortality to reduce the frog population size to below NT however, persistence of the pathogen would then require a delicate balance between mortality due to chytridiomycosis and replenishment of the host population through reproduction. Investigating this requires additional assumptions about frog reproduction and mortality a number of variants were evaluated.

Unstructured host model.

In the simplest version, there is no stage structure in the host population, and frog reproduction occurs in a discrete pulse each year. With an unstructured host population, persistence of an infected frog population for even a decade is possible for only a narrow range of parameters (Fig. 3A) with relatively low self-reinfection rates and intermediate zoospore encounter rates. With very low encounter rates, Bd goes extinct, and for higher encounter rates, Bd rapidly drives the frogs toward extinction. For parameters in the persistent region, within a year the fungal load on a fraction of the individuals grows exponentially (e.g., black line in Fig. 3B) until Smax is reached, at which point those frogs die, whereas the fungal loads on another fraction of the individuals (e.g., red, blue, and green lines in Fig. 3B) fluctuate at low levels, and in many cases individuals temporarily become uninfected and are subsequently reinfected.

Probability of persistence, and sample within-season trajectories for three different variants of the stochastic model. (A, C, and E) Probability of frogs and Bd persisting for at least 10 years as a function of reinfection rate, f, and zoospore encounter rate, γ. Shown are the fractions of 100 runs for each combination of parameters that persist for at least 10 years (red, 100% of runs persist blue, 0% of runs persist). All runs are initialized with a single infected frog in an otherwise uninfected frog population at its carrying capacity and no zoospores in the zoospore pool (Z = 0). In all of the models, the frog population can be rapidly driven extinct at high values of the zoospore encounter rate (high γ). (B, D, and F) Examples of within-season dynamics showing the dynamics of the number of sporangia on individual frogs. The colored lines are highlighted examples of trajectories of sporangia on individual frogs. (A and B) Unstructured host model, with R = 4, K = 100 frogs, θF = 0.9, θZ = 0.1, V = 1 unit volume, ν = 0.1, μ = 1 day −1 , η = 17.5 zoospores · day −1 , and σ = 0.2 day −1 . In B, f = 0.15, γ = 1 × 10 −6 unit volume · day −1 . (C and D) Model with external source of zoospores. All parameters are as in A, with εZ = 1,000 zoospores. In D, f = 0.1, γ = 1 × 10 −4 unit volume · day −1 . (E and F) Model with long-lived tadpole stage. In F, f = 0.05 for both tadpoles and adults, γadult = 1 × 10 −6 unit volume · day −1 , γtadpole = 100 · γadult, νadult = νtadpole = 0.1, Smax_adult = Smax_tadpole = 10,000 sporangia, R = 40 tadpoles, θadult = 0.9, θtadpole = 0.2, θZ = 0.1, m = 0.5, V = 1 unit volume, γ = 0.01 unit volume · day −1 , μ = 1 day −1 , η = 17.5 zoospores · day −1 , and σ = 0.2 day −1 . Stars in A, C, and E indicate parameter values used for simulations in B, D, and F, respectively.

Model with an external source of zoospores.

Long-term persistence is a more likely outcome if there is some mechanism present to keep the pathogen from going extinct during the troughs of zoospore density. An external source of zoospores, from either an environmental reservoir for Bd (an environmental reservoir for Bd has not yet been found, although this remains a possibility) or a more resistant alternative amphibian host that contributes a constant input of zoospores into the zoospore pool, can result in persistence of an infected frog population over a wide range of parameter values (Fig. 3C). For low values of the zoospore encounter rate, individual frogs can repeatedly gain and lose the infection without ever reaching the high fungal loads at which Bd causes mortality (Fig. 3D). This pattern is consistent with what we observe in persistent mountain yellow-legged frog populations.

Model with a long-lived tadpole stage.

A long-lived tadpole stage that can become infected but not succumb to infection also can promote pathogen persistence (Fig. 3E). In this model variant, we assume that both tadpoles and adults become infected by, and contribute to, the zoospore pool, but that tadpoles do not die when their Bd load reaches the maximum number of zoospores per tadpole. As such, tadpoles can act as a within-host reservoir for the pathogen, producing zoospores that can infect susceptible adults. Model results show that the pathogen can persist on a stage-structured host population across a wide range of parameters, especially when transmission rates are relatively low (Fig. 3E). In the persistent region of parameter space with low self-reinfection rates, the tadpoles continually transmit zoospores to the adult population, maintaining low to intermediate zoospore loads on adults (e.g., the red line in Fig. 3F), with no adult mortality due to Bd. Newly recruited adults rapidly become infected with low zoospore loads (e.g., the blue line in Fig. 3F).

Model with full R. Muscosa/R. Sierrae stage structure.

In a variant of the model with the full frog stage structure (17), the potential for persistence is again enhanced by the multiyear tadpole stage. Figure 4 shows three examples of the types of model dynamics observed in field populations of R. muscosa/R. sierra, where the only difference leading to the different trajectories is a change in the transmission parameter, γ. In Fig. 4A, subadults survive to recruit to the reproductive adult stage only occasionally, whereas Bd loads on adults never reach the lethal level. In the model, this occurs when zoospores encounter and successfully infect subadults at a higher rate, but subadults die of chytridiomycosis at a lower fungal load (i.e., lower Smax) than adults. In Fig. 4B, frogs and Bd remain extant for many years after arrival of Bd at a site, but subadults never recruit to the adult stage, and the apparently persistent population is actually on a slow decline to extinction. Figure 4C shows an example of the type of dynamics observed in many R. muscosa/R. sierrae populations that rapidly go extinct after the arrival of Bd. Adults and subadults reach high Bd loads and die shortly after Bd arrival. All individuals in each tadpole class die due to chytridiomycosis as soon as they reach metamorphosis, and the population is completely extinct within 2–3 years. This occurs when the adults encounter zoospores at a high rate.

Examples of dynamics of model with full-stage R. muscosa/R. sierra structure. The simulation starts with the population uninfected, and Bd invades during year 20 of the simulation. For A, the within-season dynamics are also shown for a single year (year 51). Thick lines in the within-season dynamics plots are simply trajectories of highlighted individuals for illustrative purposes. In A, γadult = 1 × 10 −6 unit volume · day −1 , γsubadult = 10 · γadult, γtadpole = 100 · γadult, Smax_adult = 10,000, Smax_subadult = Smax_tadpole = 1,000. In B. γadult = 1 × 10 −4 unit volume · day −1 , γsubadult =10 · γadult, γtadpole = 100 · γadult, Smax_adult = 10,000, and Smax_subadult = Smax_tadpole = 1,000. In C, γadult = γsubadult = γtadpole =1 × 10 −3 unit volume · day −1 , Smax_adult = Smax_subadult = Smax_tadpole = 10,000, θadult = 0.9, θsub1 = θsub2 = 0.7, θtad1 = θtad2 = θtad3 = 0.7, θZ = 0.5, m = 0.5, ωmetamorph = 0.9, pF = 0.25, R = 100, K = 100, f = 0.1, ν = 0.1, η = 17.5 zoospores · day −1 , σ = 0.2 day −1 for all frog stages, V = 1 unit volume, and μ = 1 day −1 .


C. abortus

C. abortus infects the genital tracts of ruminants (sheep, cattle, and goats) and mostly causes abortions. Infections by C. abortus can also cause stillbirths and the delivery of weak full-term newborns. C. abortus infections are the most common cause of abortions in sheep, known as ovine enzootic abortion (OEA) or enzootic abortion of ewes (EAE). The organism is usually transmitted during lambing seasons, as infected animals undergoing abortions secrete large amounts of organisms in placental tissues, uterine discharges, and in feces. In the initial stages of infection, the organism is found in the main organs of the ewe. Infection remains subclinical until the last 4 weeks of pregnancy, when abortions occur. At this time, the placenta becomes infected, while the organism is cleared from the other parts of the animal. The fetus becomes infected, leading to death and subsequent abortion. Infected ewes generally recover with their subsequent fertility unaffected. Diagnosis is made by identifying the bacteria in impression smears of placental cotlyedons, by isolation of the organism by passage in tissue culture cells, and by detection of C. abortus-specific antibodies. Infections can be treated with oral chlortetracycline if administered prior to the appearance of the second chlamydemia. However, a more effective treatment plan includes intramuscular injections of oxytetracycline. C. abortus infections are controlled by management procedures and by vaccination. Human infection due to exposure to aborting sheep has been documented and can lead to abortion or severe respiratory disease in nonpregnant individuals.

Shipping Fever Pneumonia

Shipping fever pneumonia, or undifferentiated fever, is a respiratory disease of cattle of multifactorial etiology with Mannheimia haemolytica and, less commonly, Pasteurella multocida or Histophilus somni (see Histophilosis) being the important bacterials agents involved. Shipping fever pneumonia is associated with the assembly into feedlots of large groups of calves from diverse geographic, nutritional, and genetic backgrounds. Morbidity in feeder calves often peaks within 7–10 days after assembly in a feedlot. Morbidity can approach 35%–50%, and case fatality is 5%–10% however, the level of morbidity and mortality strongly depends on the array of risk factors present in the cattle being fed.


The pathogenesis of shipping fever pneumonia involves stress factors, with or without viral infection, interacting to suppress host defense mechanisms, which allows the proliferation of commensal bacteria in the upper respiratory tract. Subsequently, these bacteria colonize the lower respiratory tract and cause a bronchopneumonia with a cranioventral distribution in the lung. Multiple stress factors are believed to contribute to suppression of host defense mechanisms. Weaning is a significant stressor, and the incidence of this disease is highest in recently weaned calves. Transportation over long distances serves as a stressor it may be associated with exhaustion, starvation, dehydration, chilling and overheating depending on weather conditions, and exposure to vehicle exhaust fumes. Additional stressors include passage through auction markets commingling, processing, and surgical procedures on arrival at the feedlot dusty environmental conditions and nutritional stress associated with a change to high-energy rations in the feedlot. The individual viral and bacterial etiologies, clinical signs, lesions, and treatment are discussed under Viral Respiratory Tract Infections in Cattle and Bacterial Pneumonia in Cattle.

Control and Prevention:

Prevention of shipping fever pneumonia should focus on reducing the stressors that contribute to development of the disease. Cattle should be assembled rapidly into groups, and new animals should not be introduced to established groups. Mixing of cattle from different sources should be avoided if possible however, in the North American beef industry, this risk factor is almost unavoidable for large intensive feedlots. Transport time should be minimized, and rest periods, with access to feed and water, should be provided during prolonged transport. Calves should ideally be weaned 2–3 wk before shipment, and surgical procedures should be performed in advance of transport however, the availability of these “preconditioned” calves is quite limited. Cattle should be processed within 48 hr after arrival at the feedlot. Adaptation to high-energy rations should be gradual, because acidosis, indigestion, and anorexia may inhibit the immune response. Vitamin and mineral deficiencies should be corrected. Dust control measures should be used.

Metaphylaxis with long-acting antibiotics such as oxytetracycline, tilmicosin, florfenicol, gamithromycin, tildipirosin, or tulathromycin has been widely adopted as a control measure given “on arrival” to cattle at high risk of developing shipping fever pneumonia. Metaphylaxis on arrival has been shown to significantly reduce morbidity and improve rate of gain and, in some cases, reduce mortality. Mass medication in feed or water is of limited value because sick animals do not eat or drink enough to achieve inhibitory blood levels of the antibiotic, and many of these oral antibiotics are poorly absorbed in ruminants.

On arrival, processing usually involves administration of modified-live vaccines for viral antigens and for bacterial components of shipping fever pneumonia. Because most cases of pneumonia occur during the first 2 wk after arrival, these on-arrival vaccines may not have adequate time to stimulate immunity. When possible, vaccinations for the viral and bacterial components of shipping fever pneumonia should be given 2–3 wk before transport or earlier and can be repeated on entry to the feedlot.

Gene expression varies within and between enzootic and epizootic lineages of Batrachochytrium dendrobatidis (Bd) in the Americas

In: Fungal Biology , Vol. 124, No. 1, 01.2020, p. 34-43.

Research output : Contribution to journal › Article




T1 - Gene expression varies within and between enzootic and epizootic lineages of Batrachochytrium dendrobatidis (Bd) in the Americas

N2 - While much research focus is paid to hypervirulent fungal lineages during emerging infectious disease outbreaks, examining enzootic pathogen isolates can be equally fruitful in delineating infection dynamics and determining pathogenesis. The fungal pathogen of amphibians, Batrachochytrium dendrobatidis (Bd), exhibits markedly different patterns of disease in natural populations, where it has caused massive amphibian declines in some regions, yet persists enzootically in others. Here we compare in vitro gene expression profiles of a panel of Bd isolates representing both the enzootic Bd-Brazil lineage, and the more recently diverged, panzootic lineage, Bd-GPL. We document significantly different lineage-specific and intralineage gene expression patterns, with Bd-Brazil upregulating genes with aspartic-type peptidase activity, and Bd-GPL upregulating CBM18 chitin-binding genes, among others. We also find pronounced intralineage variation in membrane integrity and transmembrane transport ability within our Bd-GPL isolates. Finally, we highlight unexpectedly divergent expression profiles in sympatric panzootic isolates, underscoring microgeographic functional variation in a largely clonal lineage. This variation in gene expression likely plays an important role in the relative pathogenesis and host range of Bd-Brazil and Bd-GPL isolates. Together, our results demonstrate that functional genomics approaches can provide information relevant to studies of virulence evolution within the Bd clade.

AB - While much research focus is paid to hypervirulent fungal lineages during emerging infectious disease outbreaks, examining enzootic pathogen isolates can be equally fruitful in delineating infection dynamics and determining pathogenesis. The fungal pathogen of amphibians, Batrachochytrium dendrobatidis (Bd), exhibits markedly different patterns of disease in natural populations, where it has caused massive amphibian declines in some regions, yet persists enzootically in others. Here we compare in vitro gene expression profiles of a panel of Bd isolates representing both the enzootic Bd-Brazil lineage, and the more recently diverged, panzootic lineage, Bd-GPL. We document significantly different lineage-specific and intralineage gene expression patterns, with Bd-Brazil upregulating genes with aspartic-type peptidase activity, and Bd-GPL upregulating CBM18 chitin-binding genes, among others. We also find pronounced intralineage variation in membrane integrity and transmembrane transport ability within our Bd-GPL isolates. Finally, we highlight unexpectedly divergent expression profiles in sympatric panzootic isolates, underscoring microgeographic functional variation in a largely clonal lineage. This variation in gene expression likely plays an important role in the relative pathogenesis and host range of Bd-Brazil and Bd-GPL isolates. Together, our results demonstrate that functional genomics approaches can provide information relevant to studies of virulence evolution within the Bd clade.

1748, in the meaning defined above

borrowed from French épizootique "pertaining to a disease affecting many animals at once," coinage modeled on épidemique epidemic entry 1, from Greek epi- epi- + zôion "animal" + French -otique -otic entry 1 — more at zoo-

Note: The Oxford English Dictionary and Trésor de la Langue Française both treat épizootique as a derivative of épizootie, but the noun is attested later and is more likely a back-formation from the adjective, on the model of épidémique : épidémie.


Contagion and infection by far play the biggest role in the dissemination and spread of epizootic and panzootic diseases. These include virulent (ex. Cattle Plague), septic (can be caused in the change in food quality), parasitic (ex. Scabies), and miasmatic infections (ex. Typhoid Fever). Many claim that an accidental morbific cause, which infects a great number of animals which ceases activity after a prolonged time period. [1]

Certain factors come into play in the spread of certain panzootic diseases, as can be seen with Batrachochytrium dendrobatidis. This infection seems to be sensitive to external conditions, particularly the environments temperature and moisture. These factors leads to limitations on where the diseases can thrive, acting almost as its ‘climate niche’. [2]

Persistence of H5N1 Avian Influenza Edit

Influenza A virus subtype H5N1, the highly pathogenic strain of influenza, was first detected in the goose population of Guangdong, China in 1996. [3]

In February 2004, avian influenza virus was detected in birds in Vietnam, increasing fears of the emergence of new variant strains. It is feared that if the avian influenza virus combines with a human influenza virus (in a bird or a human), the new subtype created could be both highly contagious and highly lethal.

In October 2005, cases of the avian flu (the deadly strain H5N1) were identified in Turkey. EU Health Commissioner Markos Kyprianou said: "We have received now confirmation that the virus found in Turkey is an avian flu H5N1 virus. There is a direct relationship with viruses found in Russia, Mongolia and China." Cases of bird flu were also identified shortly thereafter in Romania, and then Greece. Possible cases of the virus have also been found in Croatia, Bulgaria and in the United Kingdom. [4] However, by the end of October only 67 people had died as a result of H5N1 which was atypical of previous influenza pandemics.

The enzooicity of H5N1 in birds, poultry in particular, has led to a major panzootic. As of 2012, there was an estimated 400 million birds killed from infection of the H5N1 strain of influenza. Studies have shown that H5N1 is very well adapted to domestic duck and geese, making them key in controlling the H5N1 strain and preventing future panzootic events. [3]

Presently, the highly pathogenic Asian strain of Avian Influenza is still continuing to kill poultry and wild birds alike on panzootic scales. The persistence of such a pathogen in the environment would only lead to a further continuation of panzootic scale eliminations of birds. To try to control this, scientists did research involving the shed feathers of domestic ducks to test the prevalence of H5N1. Although viral persistence was notably found within drinking water and feces, the feathers exhibited the most remaining H5N1 strain for up to 160 days. [5] The persistence exhibited through the feathers indicates the potential for environmental contamination of not only H5N1, but also other untested viruses.

White Nose Syndrome in Bats Edit

White Nose Syndrome (WNS) is a rapidly spreading fungal infection responsible for killing millions of bats within the past 9 years. [6] Geomyces-destructans is the causative fungal agent of the characteristic skin lesions seen on the exposed skin, particularly on wings and nose, and in the hair follicles of affected bats. WNS has only recently been discovered, in Howe's Cave, New York during the winter of 2006-2007, [7] but affects 25% of the hibernating species. [8] Six species of bats have been fatally effected by this panzootic big brown bat, small-footed bat, little brown bat, northern long-eared bat, Indiana bat, and tricolored bat, and current bat population surveys suggest a 2-year population decline in excess of 75%. [9] The geographical range of WNS has spread throughout 33 states, and 4 Canadian provinces. [8] [9]

The mechanism of how the infection on the skin kills bats is unclear. [8] However, the outward cause of mortality of the infected bats depends on the stage and severity of the bat. Infected bats commonly die from starvation over winter, and can suffer from lasting damage to the wing membranes and impair summer foraging if survived the winter. [10] One of the most abundant bat species in eastern North America, the little brown bat (Myotis lucifugus), could disappear from this region within 16 years. [10]

Severely infected bats emerge prematurely from hibernation, and if they survive long enough and enter a different hibernaculum, the likelihood of transmission is probably high, because they presumably carry a large load of fungal spores. [8] Transmission of the infection is either physically from bat-to-bat contact, or from and hibernaculum-to-bat, through the exposure to spores of Geomyces- destructans that were present on a roosting substrate. [8]

Newcastle Disease in Pigeons Edit

Newcastle disease is a contagious bird disease affecting many domestic and wild avian species. [11] The disease is contagious through immediate contact between healthy birds and the bodily discharges of infected birds. This includes transmission through droppings, secretions from the nose, mouth and eyes. Newcastle disease spreads quickly among birds kept in captivity, such as commercially raised chickens. [12] Symptoms include sneezing, gasping for air, nasal discharge, coughing, greenish and watery diarrhea, nervousness, depression, muscular tremors, drooping wings, twisting of head and neck, circling, complete paralysis, partial to complete drop in egg production and thin-shelled eggs, swelling of the tissues around the eyes and in the neck, and sudden death. [12]

Newcastle disease was first identified in Java, Indonesia, in 1926, and in 1927, in Newcastle upon Tyne, England (whence it got its name). However, it may have been prevalent as early as 1898, when a disease wiped out all the domestic fowl in northwest Scotland. [13] Its effects are most notable in domestic poultry due to their high susceptibility and the potential for severe impacts of an epizootic on the poultry industries. It is endemic to many countries. The emergence and spread of new genotypes across the world represents a significant threat to poultry. This suggests that the disease is continuously evolving, leading to more diversity (Miller et al., 2009). [14]

Unfortunately, little has been done to comprehend the procedure and advancement of new genotypes (Alexander et al., 2012). [14] Recent vNDV have been characterized as isolates and offer evidence which proposes an emergence of a fifth panzootic initiated by highly related vNDV isolates from Indonesia, Israel and Pakistan. [15] These virus strains are related to older strains from wild birds. This suggests that unknown reservoirs harbor new vNDV isolates capable of additional panzootics.

No treatment for NDV exists, but the use of prophylactic vaccines [16] and sanitary measures reduces the likelihood of outbreaks.

Chytrid Fungus in Amphibian Populations Edit

Chytridiomycosis caused by a chytrid fungus is a deadly fungal disease that has wiped out 30 amphibian species across the globe and has sent overall amphibian populations in decline. The fungus Batrachochytrium dendrobatidis can be found on every continent with fertile soil and has contributed to the loss of some species of frogs and salamanders. [17] In fact, it is estimated that 287 species of amphibians are infected with this disease in over 35 countries. [18] These countries tend to have varied or tropical climates like those found in Central America, South America, and Africa with optimal climate conditions ranging from 17 degree Celsius to 23 degrees Celsius for the fungus to thrive. [17]

The first reported instance of the chytrid fungus was in 1998 which was found on the skin of amphibians. [17] Since amphibians absorb essential nutrients through their skin, the fungus attaches itself to the amphibian, suffocates the amphibian, and enters the blood stream of the organism. Some symptoms that are prevalent in affected species include lethargy and loss of equilibrium and begin to die 21 days after infection. [17] Frogs that have died and are examined show high density of the fungal spores in keratinized areas of the body such as the pelvis, mouth, and underbelly.

The fungus is spread through the transportation of amphibious species by humans. Infected amphibians that have escaped or are released into the wild can carry the fungus and therefore invade the surrounding habitats of local species that are not immune to the disease. Species like the American Bullfrog and African Clawed Frog can carry this disease without experiencing symptoms or death these kinds of species are usually to blame for the spread of the disease in undeveloped habitats. [19]

Some characteristics of amphibians that are more likely to be susceptible to the disease are the lack of various developed microbiota that live and breed on the dermis of the species as well the underdeveloped immune system in specific amphibians. [17] Species that tend to breed in flowing water which washes away the microbiota from the skin of amphibians are more likely to become infected. Organizations across the world have tried to implement rules and regulations for the transportation of amphibians across borders to prevent the continued decline of amphibians however progress has been slow. [17] To add to the slow progress, the only cure that exists for chytrid fungus is found within laboratories for amphibians in captivity. Because of this, there is no known way for eradicating the disease in the wild.

Natural enzootic vectors of Venezuelan equine encephalitis virus, Magdalena Valley, Colombia. (Research).

To characterize the transmission cycle of enzootic Venezuelan equine encephalitis virus (VEEV) strains believed to represent an epizootic progenitor, we identified natural vectors in a sylvatic focus in the middle Magdalena Valley of Colombia. Hamster-baited traps were placed into an active forest focus, and mosquitoes collected from each trap in which a hamster became infected were sorted by species and assayed for virus. In 18 cases, a single, initial, high-titered mosquito pool representing the vector species was identified. These vectors included Culex (Melanoconion) vomerifer (11 transmission events), Cx. (Mel.) pedroi (5 transmissions) and Cx. (Mel.) adamesi (2 transmissions). These results extend the number of proven enzootic VEEV vectors to 7, all of which are members of the Spissipes section of the subgenus Melanoconion. Our findings contrast with previous studies, which have indicated that a single species usually serves as the principal enzootic VEEV vector at a given location.

Venezuelan equine encephalitis (VEE) is an emerging oonotic arboviral disease that affects equines and humans in the Americas (1). Venezuelan equine encephalitis virus (VEEV) has caused sporadic outbreaks since the early part of the 20th century, with some epidemics affecting > 100,000 persons. For many years, the source of the epizootic/epidemic VEEV strains belonging to subtypes IAB and IC viruses remained unknown. After antigenically related but distinct, equine-avirulent, enzootic strains of VEEV were isolated in the 1960s, researchers hypothesized that epizootic/epidemic strains evolve from enzootic VEEV progenitors (2). The first genetic evidence supporting this hypothesis came from RNA fingerprinting studies that indicated a close relationship between subtype ID-enzootic VEEV strains from Colombia and epizootic/epidemic isolates belonging to subtype IC (3). Later, sequencing (4) and phylogenetic (5,6) studies also supported the evolution of the epizootic/epidemic serotype IAB and IC strains from enzootic ID VEEV progenitors. Recently, comprehensive phylogenetic analyses have indicated that the epizootic/epidemic strains evolved independently on at least three occasions from a single lineage of ID VEEV that circulates in eastern and central Colombia, western Venezuela, and northern Peru (7-10). Other ID-like VEEV lineages that occur in Panama, Amazonian Peru, southwestern Colombia, coastal Ecuador, north-central Venezuela, and Florida have not generated any of the epizootic/epidemic strains sequenced (10-12).

Enzootic VEEV (subtypes ID-IF, II-VI) circulate nearly continuously in sylvatic or swamp habitats in various tropical and subtropical locations in the New World (1,13). These viruses generally use small mammals as their reservoir hosts and are transmitted by mosquitoes. Enzootic mosquito vectors have been identified for four VEEV variants: 1) Culex (Melanoconion) portesi transmits Mucambo virus (VEE complex subtype IIIA) in Trinidad (14), 2) Cx. (Mel.) cedecei transmits Everglades virus (VEE complex subtype II) in southern Florida (15), 3) Cx. (Mel.) aikenii sensu lato (ocossa and panocossa) transmits subtype ID VEEV in Panama (16,17), and 4) Cx. (Mel.) taeniopus (formerly opisthopus) is the primary enzootic vector of subtype IE VEEV in Guatemala (18). More than 70% of enzootic field isolations have come from the subgenus Melanoconion, suggesting that these mosquitoes are the principal vectors of most or all enzootic VEE complex strains (17).

The infrequency of VEE emergence is probably determined by the infrequent, simultaneous occurrence in time and space of viral mutations that mediate host range changes, combined with ecologic and epidemiologic conditions that permit efficient amplification (1). To understand the mechanisms of VEE emergence from enzootic progenitors in Colombia and Venezuela, we are studying the hosts in which epizootic mutations may occur and in which the selection of epizootic strains may follow. However, the vector and reservoir hosts of the particular subtype ID VEEV lineage implicated in epizootic emergence have not been identified. Using an efficient system of vector identification employing hamster baited traps, we identified Cx. (Mel.) vomerifer, Cx. (Mel.) pedroi, and Cx. (Mel.) adamesi as natural enzootic vectors in an active focus of subtype ID VEEV in the middle Magdalena Valley of Colombia.

The study was carried out from 1999 to 2000 in the Monte San Miguel Forest in the middle Magdalena Valley of Colombia (6[degrees] 23' 30"N 74[degrees] 21' 41' W 50 m elevation). This is a lowland tropical rainforest surrounded by cattle ranches created by deforestation. Mean minimum and maximum daily temperatures are 23[degrees]C and 33[degrees]C (overall mean of 29[degrees]C), respectively, and annual rainfall averages 2,700 mm. Mean relative humidity is 80%. Generally, the peaks of the rainy seasons occur in April-May and October-November. Numerous previous isolations of subtype ID VEEV from sentinel hamsters (9) indicate that this forest site is a stable enzootic focus.

Hamster-baited traps were used for detection of natural VEEV vectors. These traps were a version of the Trinidad No. 10 trap (19) with the following modifications: 1) the metal can comprising the trap opening was replaced by a polyvinyl chloride pipe, 10 cm in diameter 2) the cylindrical animal cage was enlarged to 11 cm in diameter and 12 cm in height 3) the roof was constructed from plexiglass and 4) the opening for mosquito aspiration was a simple buttonhole sewn into the polyester collection net (Figure). The traps were baited with adult golden Syrian hamsters obtained from a colony maintained at the Instituto Nacional de Salud in Bogota. Baited traps were suspended approximately 1.5 m above the ground and placed in transects at 10-m intervals. Carrots and rat chow were provided for food and water. The traps were checked each morning between 0600 and 0800 h, and some were also checked in the evening between 1700 and 1900 h. Mosquitoes were removed from the traps by using an aspirator, and the daily or semi-daily collections from each trap were frozen as a single pool in a plastic bottle immersed in liquid nitrogen vapor. When hamsters within the traps became moribund or died, serum samples were obtained by cardiac puncture or their hearts were dissected aseptically and frozen for virus isolation.

Detection of Natural Transmission to Hamsters

To confirm VEEV infection in dead or moribund hamsters, virus was isolated from a 10% heart tissue suspension in Eagle's minimal essential medium (MEM), supplemented with 20% fetal bovine serum (FBS) and antibiotics. The suspension was prepared in a Ten Broeck tissue grinder and centrifuged at 15,000 x g for 5 min, 200 [micro]L of the supernatant was added to a 25-[cm.sup.2] flask containing a monolayer of Vero cells and adsorbed for 1 h at 37[degrees]C 6 mL of additional MEM containing 2% FBS was then added. Cultures were incubated at 37[degrees]C for 5 days or until cytopathic effects were evident.

Mosquito pools from traps in which hamster infection with VEEV was confirmed were assayed for infectious virus. Pools containing 1-40 individuals of each mosquito species were triturated with a Minibeadbeater (BioSpec Products, Inc., Bartlesville, OK) or a Ten Broeck tissue grinder containing 1.0 mL of MEM supplemented with 20% FBS, penicillin, streptomycin, and amphotericin B. The triturated pool was centrifuged for 5 min at 15,000 x g, and 200 [micro]l of the supernatant was added to a 10-mL plastic tube or a 25-[cm.sup.2] cell culture dish containing a monolayer of Vero cells and 2-5 mL of MEM. Cultures were monitored for cytopathic effects for 5 days.

Genetic and Antigenic Characterization of VEEV Isolates

Viruses isolated from hamster heart tissue suspensions and mosquito pools were characterized antigenically by using immunofluorescence of infected cells and a panel of monoclonal antibodies described previously (20). Subtype ID VEEV isolates were further characterized genetically by reverse transcription-polymerase chain reaction (PCR) amplification of an 856-nucleotide portion of the PE2 (sometimes called p62) envelope glycoprotein precursor gene as described previously (8), followed by single-stranded conformation polymorphism (SSCP) or sequence analysis (9). For SSCP analysis, PCR products were purified on agarose gels by using the QIAquick Gel Extraction Kit (QIAGEN, Valencia, CA). A 2-[micro]l volume of the PCR amplicon DNA suspension was mixed with 8 [micro]l of SSCP loading buffer (95% formamide, 0.05% bromophenol blue, and 0.05% xylene cyanol). The DNA was heated to 95[degrees]C for 5 minutes, rapidly cooled on ice, loaded onto an 8% polyacrylamide gel, and underwent electrophoresis in 1X Tris-borate EDTA buffer at room temperature for 20 h at 8 mA. Single-stranded DNA products were visualized by using silver staining (21). SSCP patterns were compared by measuring the migration of single-stranded DNA of the various isolates in comparison to one another and to a standard DNA ladder.

For vector identification studies, 87 hamsters were exposed in traps within the Monte San Miguel Forest for 5-7 days. Of these, 38 became moribund or died and were processed for virus isolation. VEEV was isolated from 37 hamsters, and the mosquito collections from the corresponding traps were assayed for virus.

In 18 of the traps yielding infected hamsters, a vector species was identified by using the following criteria: 1) the hamster died at least 24 h after the collection of the presumed vector, consistent with the incubation time of VEEV in hamsters (22) 2) during the first day in which infected mosquitoes were collected from the trap, only one species pool had a high titer (>5.0 [log.sub.10] PFU/pool) consistent with an infectious mosquito, as determined by previous experimental studies of enzootic VEEV vectors (18,23-26) 3) the remaining pools, from the first day in which infected mosquitoes were collected, were uninfected, or had low titers (<5.0 log) shown previously to be inconsistent with an infectious mosquito (18,23-26) 4) the mosquito collections on the days subsequent to that of the vector collection were mostly infected, reflecting hamster viremia and the ingestion of infectious blood by mosquitoes biting [greater than or equal to] 12 h after the transmission event and 5) virus isolates from the hamster and corresponding vector were indistinguishable antigenically and genetically with SSCP analysis, sequencing, or both. In 18/37 infected hamster events studied, these criteria were fulfilled, vector was identified unambiguously. Typical data for one of these transmission events (hamster 164) is shown in Table 1. In this example, transmission by Cx. vomerifer occurred [less than or equal to] 24 h after exposure of the trap, and the vector pool had a titer of 5.8 [log.sub.10] PFU/ pool. The other two infected pools from day 2, Cx. pedroi and Aedes serratus, had log titers [less than or equal to] 3.3, indicating that they were not capable of transmission. These pools presumably contained one or more mosquitoes that engorged on the hamster after viremia began, probably just before the daily trap collection. On the next day, all mosquito pools contained infectious virus in their midguts, representing viremic hamster blood ingested by mosquitoes within the trap.

A total of 18 transmission events were characterized as described above. The most common interval of collection of the identified vector was 24--48 h after exposure, reflecting a very high level of enzootic VEEV transmission in the Monte San Miguel Forest. Cx. vomerifer was implicated in 11 of these events, Cx. pedroi in 5, and Cx. adamesi in 2 transmissions (Table 2). The minimum infection/transmission rate for the mosquitoes we collected could not be determined directly because we did not identify the mosquito collections for traps where transmission to the hamster did not occur. However, rates on the order of 1/200-1/1000 can be estimated for these three vector species if the species composition is assumed to be similar in traps where transmission did not occur. Even if this assumption is incorrect, the error in this estimate should not be more than twofold because VEEV transmission occurred in most traps.

Use of Hamster-Baited Traps for Arbovirus Vector Identification

Traditional criteria for arthropod vector identification include the following: 1) demonstration of feeding or other effective contact with pathogen's host 2) association in time and space of the vector and pathogen 3) repeated demonstration of natural infection of the vector, and 4) experimental transmission of the pathogen by the vector (27). Infection rates for arbovirus vectors tend to be relatively low, usually <1%. Therefore, fulfillment of these criteria for arbovirus vectors usually relies on the capture of large numbers of arthropods for virus isolation, followed by experimental laboratory transmission studies to ensure that species found infected in nature are competent vectors. Although this strategy is the most comprehensive and unbiased, it is extremely costly and time consuming, accounting for the relative paucity of information on natural vectors of many arboviruses. Some studies of VEEV vectors have also relied on oral infection from experimentally infected hamsters with viremia levels of very high titer, on the order of 8 [log.sub.10] PFU/mL (28,29), a titer at least 100-1,000 times greater than that generated by experimentally infected rodent reservoir hosts (30,31), equines (13,30,32), or naturally infected humans (8,33) (Some studies of equine viremia have yielded titers of >[10.sup.8] suckling mouse intracerebral 50% lethal doses, but this method for quantifying VEEV titers is 100- to 1000-fold more sensitive than PFU [30,34]). Results from these studies are therefore inconclusive regarding natural transmission potential.

Other investigators have streamlined the vector identification process by collecting suspected vectors and sorting them according to species, then exposing single-species pools to naive animals in a field or laboratory setting to detect transmission (16,18). We have taken this approach one step further by combining collection and transmission detection using hamster-baited traps. This method simplifies the vector identification process in several ways: 1) Hamster-baited traps attract and capture only arthropod species that are attracted to small mammals, the natural reservoir hosts of the enzootic VEEV (Proechimys spp. spiny rats in the case of subtype ID VEEV circulating in this focus [35]), minimizing collection and mosquito processing efforts. 2) Arthropod collections from traps where no transmission occurs do not need to be sorted, greatly reducing a laborious step in the vector identification process. 3) Only a small number of arthropod pools must be tested for virus, eliminating much of the cost, labor, and biosafety hazard associated with traditional vector identification approaches. In addition, the hamster-baited traps can serve as sentinels for detection of active virus circulation in a forest and reveal the presence of other viruses in a focus. However, unlike other sentinel enclosures that allow arthropods to escape after biting a viremic bait animal and thereby initiate artificial amplification, the hamster-baited traps capture most of the arthropods that bite the viremic host and prevent most or all artificial amplification. A similar strategy for detecting transmission of western equine encephalitis and St. Louis encephalitis viruses to chickens in baited traps was described by Reeves et al. (36).

Using these hamster-baited traps alone, we were not able to measure directly the capture efficiency of our traps. However, in the case of five infected hamsters, the lack of any collections with a single or few high titer mosquito species pools on the day preceding total infection of collected mosquitoes indicates that the arthropod responsible for transmission may have escaped. In other cases, two or more mosquito pools collected on the first day virus was detected had titers consistent with infectious vectors, precluding vector identification. We are currently experimenting with funnel-shaped openings to reduce the frequency of vector escape from this trap design. As with any passive trap design, a compromise between ease of vector entry and frequency of escape must be sought to maximize collections.

Enzootic Vectors of Venezuelan Equine Encephalitis Complex Viruses

Previous studies of VEE complex enzootic transmission have each identified a single, principal mosquito species in a given geographic region. All of these species, including Cx. portesi (14), Cx. cedecei (15), Cx. aikenii sensu lato (ocossa and panocossa) (16,17), and Cx. taeniopus (18) are members of the Spissipes section of the subgenus Melanoconion within the genus Culex (37). Previous studies of enzootic VEEV transmission in the Catatumbo region of northeastern Colombia also suggested that Cx. pedroi might be the principal vector, based on abundance in active foci (38). Cx. vomerifer from Iquitos, Peru, also has been shown to be susceptible to infection by several strains of VEEV (28), but was only tested after mosquitoes ingested 8 [log.sub.10] PFU/mL from viremic hamsters, a viremia titer at least 100 times greater than that generated by experimentally infected rodent reservoir hosts (30,31). Our findings of at least three enzootic vectors of subtype ID VEEV in Colombia contrast with the findings of all previous studies of enzootic VEEV vectors, which suggested that enzootic VEEV strains are each adapted to a single, principal vector species (13,18,39-41). In Colombia, subtype ID VEEV appears to utilize efficiently both Cx. vomerifer and Cx. pedroi in the Magdalena Valley. Cx. adamesi, which is usually less abundant in the Monte San Miguel Forest, appears to serve as a secondary vector.

All three of the mosquito species that we identified as VEEV vectors are members of the Spissipes section of the subgenus Culex (Melanoconion), bringing the total to seven confirmed vectors within this section of closely related mosquitoes. The genetic or ecologic basis for the exclusive use of these mosquitoes by enzootic VEE complex viruses deserves further study. Hypotheses to explain this phenomenon include possible shared, derived characteristics of the Spissipes section, such as particularly high susceptibility to infection by enzootic VEE complex viruses, a particularly high degree of association with the Proechimys spp. (35) and other small mammalian reservoir hosts (13), or both. Mosquito longevity and population sizes in habitats that support large populations of reservoir hosts may also favor transmission by members of the Spissipes section (35).

Role of Enzootic Vectors in VEEV Emergence and Disappearance

Identification of the principal enzootic vectors (Cx. vomerifer and Cx. pedroi) of subtype ID VEEV strains believed to be closely related to epizootic progenitors will allow us to assess the role of these mosquitoes in the generation of mutations that mediate VEE emergence by enhancing equine viremia and infection of epizootic mosquito vectors such as Ochlerotatus taeniorhynchus. The hypothesis that epizootic VEEV is not recovered from sylvatic foci because these strains lose their fitness for the enzootic vectors (25) can also be tested in the two principal vectors that we identified.

We thank Marco Fidel Suarez and Eutimio Guerra for excellent technical assistance.

This research was supported by grants AI39800 and AI48807 from the National Institutes of Health, and by Colciencias grant 210404-758-98.

(1.) Weaver SC. Recurrent emergence of Venezuelan equine encephalomyelitis. In: Scheld WM, Hughes J, editors. Emerging infections I. Washington, D.C.: ASM Press 1998. p. 27-42.

(2.) Johnson KM, Martin DH. Venezuelan equine encephalitis. Adv Vet Sci Comp Med 197418:79-116.

(3.) Rico-Hesse R, Roehrig JT, Trent DW, Dickerman RW. Genetic variation of Venezuelan equine encephalitis virus strains of the ID variety in Colombia. Am J Trop Med Hyg 198838:195-204.

(4.) Kinney RM, Tsuchiya KR, Sneider JM, Trent DW. Genetic evidence that epizootic Venezuelan equine encephalitis (VEE) viruses may have evolved from enzootic VEE subtype I-D virus. Virology 1992191:569-80.

(5.) Rico-Hesse R, Weaver SC, de Siger J, Medina G, Salas RA. Emergence of a new epidemic/epizootic Venezuelan equine encephalitis virus in South America. Proc Natl Acad Sci U S A 199592:5278-81.

(6.) Weaver SC, Bellew LA, Rico-Hesse R. Phylogenetic analysis of alphaviruses in the Venezuelan equine encephalitis complex and identification of the source of epizootic viruses. Virology 1992191:282-90.

(7.) Wang E, Barrera R, Boshell J, Ferro C, Freier JE, Navarro JC, et al. Genetic and phenotypic changes accompanying the emergence of epizootic subtype IC Venezuelan equine encephalitis viruses from an enzootic subtype ID progenitor. J Virol 199973:4266-71.

(8.) Weaver SC, Salas R, Rico-Hesse R, Ludwig GV, Oberste MS, Boshell J, et al. Re-emergence of epidemic Venezuelan equine encephalomyelitis in South America. Lancet 1996348:436-40.

(9.) Moncayo AC, Medina GM, Kalvatchev Z, Brault AC, Barrera R, Boshell J, et al. Genetic diversity and relationships among Venezuelan equine encephalitis virus field isolates from Colombia and Venezuela. Am J Trop Med Hyg 200165:738-46.

(10.) Powers AM, Oberste MS, Brault AC, Rico-Hesse R, Schmura SM, Smith JF, et al. Repeated emergence of epidemic/epizootic Venezuelan equine encephalitis from a single genotype of enzootic subtype ID virus. J Virol 199771:6697-705.

(11.) Salas RA, Garcia CZ, Liria J, Barrera R, Navarro JC, Medina G, et al. Ecological studies of enzootic Venezuelan equine encephalitis in north-central Venezuela, 1997-1998. Am J Trop Med Hyg 200164:84-92.

(12.) Brault AC, Powers AM, Holmes EC, Woelk CH, Weaver SC. Positively charged amino acid substitutions in the E2 envelope glycoprotein are associated with the emergence of Venezuelan equine encephalitis virus. J Virol 200276:1718-30.

(13.) Walton TE, Grayson MA. Venezuelan equine encephalomyelitis. In: Monath TP, editor. The arboviruses: epidemiology and ecology, vol. IV. Boca Raton (FL): CRC Press 1988. p. 203-31.

(14.) Aitken THG. Habits of some mosquito hosts of VEE (Mucambo) virus from northeastern South America, including Trinidad. In: Proceedings of workshop-symposium on Venezuelan encephalitis virus. Washington: Pan American Health Organization Scientific Publ. 2431972. p. 254-6.

(15.) Chamberlain RW, Sudia WD, Coleman PH, Work TH. Venezuelan equine encephalitis virus from south Florida. Science 1964145:272-4.

(16.) Galindo P, Grayson MA. Culex (Melanoconion) aikenii: natural vector in Panama of endemic Venezuelan encephalitis. Science 1971172:594-5.

(17.) Galindo P. Endemic vectors of Venezuelan encephalitis. In: Proceedings of workshop-symposium on Venezuelan encephalitis virus,. Washington: Pan American Health Organization Scientific Publ. 243 1972. p. 24953.

(18.) Cupp EW, Scherer WF, Ordonez JV. Transmission of Venezuelan encephalitis virus by naturally infected Culex (Melanoconion) opisthopus. Am J Trop Med Hyg 197928:1060-3.

(19.) Davies JB. A small mosquito trap for use with animal or carbon dioxide baits. Mosquito News 197131:441-3.

(20.) Roehrig JT, Bolin RA. Monoclonal antibodies capable of distinguishing epizootic from enzootic varieties of subtype I Venezuelan equine encephalitis viruses in a rapid indirect immunofluorescence assay. J Clin Microbiol 199735:1887-90.

(21.) Black WC, Vanlandingham DL, Sweeney WP, Wasieloski LP, Calisher CH, Beaty BJ. Typing of LaCrosse, snowshoe hare, and Tahyna viruses by analyses of single-strand conformation polymorphisms of the small RNA segments. J Clin Microbiol 199533:3179-82.

(22.) Scherer WF, Dickerman RW, Chia CW, Ventura A, Moorhouse A, Geiger R, et al. Venezuelan equine encephalitis virus in Veracruz, Mexico, and the use of hamsters as sentinels. Science 1963 145:274-5.

(23.) Scherer WF, Cupp EW, Lok JB, Brenner RJ, Ordonez JV. Intestinal threshold of an enzootic strain of Venezuelan equine encephalomyelitis virus in Culex (Melanoconion) taeniopus mosquitoes and its implication to vector competency and vertebrate amplifying hosts. Am J Trop Med Hyg 198130:862-9.

(24.) Weaver SC, Scherer WF, Cupp EW, Castello DA. Barriers to dissemination of Venezuelan encephalitis viruses in the Middle American enzootic vector mosquito, Culex (Melanoconion) taeniopus. Am J Trop Med Hyg 198433:953-60.

(25.) Scherer WF, Weaver SC, Taylor CA, Cupp EW. Vector incompetency: its implication in the disappearance of epizootic Venezuelan equine encephalomyelitis virus from Middle America. J Med Entomol 198623:23-9.

(26.) Weaver SC, Scherer WF, Taylor CA, Castello DA, Cupp EW. Laboratory vector competence of Culex (Melanoconion) cedecei for sympatric and allopatric Venezuelan equine encephalomyelitis viruses. Am J Trop Med Hyg 198635:619-23.

(27.) Barnett HC. The incrimination of arthropods as vectors of disease. Proceedings of the 11th Congress on Entomology, Vienna, Austria. 19602:341-5.

(28.) Turell MJ, Jones JW, Sardelis MR, Dohm DJ, Coleman RE, Watts DM, et al. Vector competence of Peruvian mosquitoes (Diptera: Culicidae) for epizootic and enzootic strains of Venezuelan equine encephalomyelitis virus. J Med Entomol 200037:835-9.

(29.) Turell MJ, Barth J, Coleman RE. Potential for Central American mosquitoes to transmit epizootic and enzootic strains of Venezuelan equine encephalitis virus. J Am Mosq Control Assoc 199915:295-8.

(30.) Wang E, Bowen RA, Medina G, Powers AM, Kang W, Chandler LM, et al. Virulence and viremia characteristics of 1992 epizootic subtype IC Venezuelan equine encephalitis viruses and closely related enzootic subtype ID strains. Am J Trop Med Hyg 200165:64-9.

(31.) Young NA, Johnson KM, Gauld LW. Viruses of the Venezuelan equine encephalomyelitis complex experimental infection of Panamanian rodents. Am J Trop Med Hyg 196918:290-6.

(32.) Walton TE, Alvarez O, Buckwalter RM, Johnson KM. Experimental infection of horses with enzootic and epizootic strains of Venezuelan equine encephalomyelitis virus. J Infect Dis 1973128:271-82.

(33.) Bowen GS, Calisher CH. Virological and serological studies of Venezuelan equine encephalomyelitis in humans. J Clin Microbiol 19764:22-7.

(34.) Martin DH, Dietz WH, Alvaerez O Jr, Johnson KM. Epidemiological significance of Venezuelan equine encephalomyelitis virus in vitro markers. Am J Trop Med Hyg 198231:561-8.

(35.) Barrera R, Ferro C, Navarro JC, Freier J, Liria J, Salas R, et al. Contrasting sylvatic foci of Venezuelan equine encephalitis virus in northern South America. Am J Trop Med Hyg 2002:67:324-34.

(36.) Reeves WC, Bellamy RE, Scrivani RP. Differentiation of encephalitis virus infection rates from transmission rates in mosquito vector populations. Am J Hyg 196173:303-15.

(37.) Sirivanakarn S. A review of the systematics and proposed scheme of internal classification of the New World subgenus Melanoconion of Culex (Diptera: Culicidae). Mosquito Systematics 198214:265-333.

(38.) Dickerman RW, Cupp EW, Groot H, Alarcon AM, Cura E, Dickerman AW, et al. Venezuelan equine encephalitis virus activity in northern Colombia during April and May 1983. Bull Pan Am Health Organ 198620:276-83.

(39.) Scherer WF, Weaver SC, Taylor CA, Cupp EW, Dickerman RW, Rubino HH. Vector competence of Culex (Melanoconion) taeniopus for allopatric and epizootic Venezuelan equine encephalomyelitis viruses. Am J Trop Med Hyg 198736:194-7.

(40.) Cupp EW, Kreutzer RD, Weaver SC. The biosystematics of Culex (Melanoconion) taeniopus sensu lato in relation to Venezuelan equine encephalomyelitis. Mosquito Systematics 198921:216-21.

(41.) Weaver SC. Vector biology in viral pathogenesis. In: Nathanson N, editor. Viral pathogenesis. New York: Lippincott-Raven 1997. p. 329-52.

Address for correspondence: Scott Weaver, Keiller 4.128, Department of Pathology, University of Texas Medical Branch, Galveston, Texas 775550609, USA fax: 409-747-2415 e-mail: [email protected]

Cristina Ferro, * Jorge Boshell, * Abelardo C. Moncayo, ([dagger]) Matra Gonzalez, * Matra L. Ahumada, * Wenli Kang, ([dagger]) and Scott C. Weaver ([dagger])

* Instituto Nacional de Salud, Bogota, Colombia and ([dagger]) University of Texas Medical Branch, Galveston, Texas, USA

Cristina Ferro is Head of the Entomology Laboratory at the National Institute of Health in Bogota, Colombia. Her research focuses on the ecology of mosquito vectors of arboviruses, especially Venezuelan equine encephalitis virus, and on sand fly vectors of leishmania and arboviruses.

Genetic diversity of enzootic isolates of vesicular stomatitis virus New Jersey.

The RNA genomes of 43 vesicular stomatitis virus (VSV) isolates of the New Jersey (NJ) serotype were T1-ribonuclease fingerprinted to compare the extent of genetic diversity of virus from regions of epizootic and enzootic disease activity. Forty of these viruses were obtained from Central America during 1982 to 1985. The other three were older isolates, including a 1970 isolate from Culex nigripalpus mosquitos in Guatemala, a 1960 bovine isolate from Panama, and a 1976 isolate from mosquitos (Mansonia indubitans) in Ecuador. The data indicate that extensive genetic diversity exists among virus isolates from this predominantly enzootic disease zone. Six distinct T1 fingerprint groups were identified for the Central American VSV NJ isolates from 1982 to 1985. The 1960 VSV NJ isolate from Panama and the 1976 isolate from Ecuador formed two additional distinct fingerprint groups. This finding is in sharp contrast to the relatively close genetic relationship existing among VSV NJ isolates obtained from predominantly epizootic disease areas of the United States and Mexico during the same period (S. T. Nichol, J. Virol. 61:1029-1036, 1987). In this previous study, RNA genome T1 fingerprint differences were observed among isolates from different epizootics however, the isolates were all clearly members of one large T1 fingerprint group. The eight T1 fingerprint groups described here for Central American and Ecuadorian viruses are distinct from those characterized earlier for virus isolates from the United States and Mexico and for the common laboratory virus strains Ogden and Hazelhurst. Despite being isolated 14 years earlier, the 1970 insect isolate from Guatemala is clearly a member of one of the 1982 to 1985 Central American virus fingerprint groups. This indicates that although virus genetic diversity in the region is extensive, under certain natural conditions particular virus genotypes can be relatively stably maintained for an extended period. The implications of these findings for the evolution of VSV NJ and epizootiology of the disease are discussed.

Watch the video: Enzootic August 2018 (February 2023).