Aquatic eutrophication promotes pathogenic infection in amphibians Pieter T. J. Johnson*†, Jonathan M. Chase‡, Katherine L. Dosch§, Richard B. Hartson§, Jackson A. Gross¶, Don J. Larson�, Daniel R. Sutherland**††, and Stephen R. Carpenter§
*Department of Ecology and Evolutionary Biology, University of Colorado, Ramaley N122, Boulder, CO 80309-0334; ‡Department of Biology, Washington University, Box 1137, St. Louis, MO 63130; §Center for Limnology, University of Wisconsin, 680 North Park Street, Madison, WI 53706-1492; ¶Southern California Coastal Water Research Project, 3535 Harbor Boulevard, Suite 110, Costa Mesa, CA 92626; �Institute of Arctic Biology, University of Alaska, P.O. Box 751403, Fairbanks, AK 99775; and **Department of Biology and River Studies Center, University of Wisconsin, 1725 State Street, La Crosse, WI 54601
Contributed by Stephen R. Carpenter, August 16, 2007 (sent for review June 18, 2007)
The widespread emergence of human and wildlife diseases has challenged ecologists to understand how large-scale agents of environmental change affect host–pathogen interactions. Acceler- ated eutrophication of aquatic ecosystems owing to nitrogen and phosphorus enrichment is a pervasive form of environmental change that has been implicated in the emergence of diseases through direct and indirect pathways. We provide experimental evidence linking eutrophication and disease in a multihost parasite system. The trematode parasite Ribeiroia ondatrae sequentially infects birds, snails, and amphibian larvae, frequently causing severe limb deformities and mortality. Eutrophication has been implicated in the emergence of this parasite, but definitive evi- dence, as well as a mechanistic understanding, have been lacking until now. We show that the effects of eutrophication cascade through the parasite life cycle to promote algal production, the density of snail hosts, and, ultimately, the intensity of infection in amphibians. Infection also negatively affected the survival of developing amphibians. Mechanistically, eutrophication promoted amphibian disease through two distinctive pathways: by increas- ing the density of infected snail hosts and by enhancing per-snail production of infectious parasites. Given forecasted increases in global eutrophication, amphibian extinctions, and similarities be- tween Ribeiroia and important human and wildlife pathogens, our results have broad epidemiological and ecological significance.
amphibian decline � emerging disease � environmental change
Emerging infections of humans and wildlife are often closelyassociated with anthropogenic alterations of the ecological and evolutionary relationships between hosts and pathogens, including climate change, biological invasions, land use change, and pollution (1–5). Owing to the complexity of pathogen– host– environment interactions, however, experimental evidence linking environmen- tal change and increased infection is often lacking, leaving the ecosystem drivers of many emerging diseases unknown (5–7). In aquatic ecosystems, one of the most profound forms of ecological change is eutrophication, which is caused by anthropogenic inputs of nitrogen (N) and/or phosphorus (P) associated with agriculture, livestock, erosion, sewage waste, and atmospheric deposition (8 – 10). Because N and P often limit primary production, their addition causes marked shifts in ecosystem conditions (8, 9, 11).
Ecological theory (12) and limited field studies (see refs. 13 and 14) suggest that, unlike many environmental stressors, eutrophica- tion will broadly enhance infection and the pathology of human and wildlife parasites. Nevertheless, experimental evidence linking nu- trient enrichment and parasitism is largely absent, and the mech- anisms through which eutrophication affects disease emergence remain poorly understood (12–15). Considering forecasted in- creases in global agricultural production and fertilizer application (10), as well as the persistence of anthropogenic P in agricultural soils and aquatic ecosystems (16), eutrophication will almost cer- tainly become an increasingly severe problem in the coming century (12). This underscores the importance of understanding the mech-
anisms linking eutrophication and host–pathogen interactions and of balancing nutrient-mediated agricultural gains with concurrent increases in disease risk (11, 14).
Here we evaluated how elevated nutrient inputs leading to eutrophication affected the transmission and pathology of Ri- beiroia ondatrae, a trematode parasite implicated in recent outbreaks of severe limb deformities in North American am- phibians. This multihost parasite, which sequentially infects freshwater snails, larval amphibians, and waterbirds [see sup- porting information (SI) Fig. 4], has been causally linked to high frequencies of malformations (10 –90%) in amphibian popula- tions, including missing limbs, extra limbs, and malformed limbs (2, 17–21) (Fig. 1). Furthermore, infection and the resulting deformities substantially reduce amphibian survival, potentially contributing to widespread population declines and extirpations. However, the ecological drivers of parasite abundance, and the reasons for the apparent increase in amphibian deformities, have remained controversial (2, 19).
We experimentally tested the hypothesis that eutrophication enhances Ribeiroia transmission and identified the mechanisms responsible for this phenomenon. Uniquely, our experiment explicitly examines how nutrient enrichment affects transmission of a disease-causing parasite among multiple hosts within a complex life cycle. We hypothesized that, by increasing algal production, eutrophication would promote parasite infection through two, potentially complementary mechanisms. First, higher resource availability will increase the population growth of susceptible snail hosts (22), leading to enhanced parasite transmission and a higher density of infected snails (12). Second, higher resource levels will reduce infected snail mortality, in- crease snail body size, and enhance host vigor (15, 23–25), promoting parasite secondary production within infected indi- viduals. Thus, a higher density of infected snails and a greater per-snail production of parasites should jointly drive an increase in amphibian infection and disease risk.
Results Nutrient additions significantly enhanced all primary response variables (Figs. 2 and 3). Except where otherwise noted, values were log10-transformed before analyses. Eutrophication pro-
Author contributions: P.T.J.J., J.M.C., J.A.G., D.R.S., and S.R.C. collected preliminary data and designed the experiments; P.T.J.J., K.L.D., and R.B.H. established the experiment, collected data, and coordinated logistics; D.J.L. and D.R.S. planned and executed amphib- ian necropsies and parasite sampling; P.T.J.J. analyzed data; and P.T.J.J., J.M.C., and S.R.C. wrote the paper.
The authors declare no conflict of interest.
Abbreviations: chl a, chlorophyll a; RM-ANOVA, repeated-measures ANOVA.
††Deceased May 26, 2006.
This article contains supporting information online at www.pnas.org/cgi/content/full/ 0707763104/DC1.
© 2007 by The National Academy of Sciences of the USA
www.pnas.org�cgi�doi�10.1073�pnas.0707763104 PNAS � October 2, 2007 � vol. 104 � no. 40 � 15781–15786
D o w
a d e d b
y g u e st
o n J
a n u a ry
1 8 , 2 0 2 2
moted growth of periphytic algae [measured as chlorophyll a (chl a)], which exhibited a monotonic increase in high nutrient mesocosms during the experiment (Fig. 2 A). We also found significant effects for time and the time-by-nutrient interaction on phytoplankton chl a [repeated-measures ANOVA (RM- ANOVA), time: Greenhouse–Geisser adjusted F[3.397,101.9] � 6.832, P � 0.0001; time-by-nutrients: Greenhouse–Geisser ad- justed F[3.397,101.9] � 3.052, P � 0.027]. However, nutrients had a significant between-subjects effect only on periphyton chl a (F[1,30] � 75.930, P � 0.0001), ref lecting the consistently stronger inf luence of nutrient enrichment on periphyton relative to phytoplankton.
Increases in periphytic algae led to enhanced egg production and a greater population biomass of herbivorous snails, regard- less of parasite input level (Fig. 2 B and C). Total snail biomass increased by nearly 50% in the high-nutrient condition but declined by 10% among low-nutrient mesocosms (Fig. 2C). Results were comparable if we used mean snail density rather than biomass. Correspondingly, snail egg masses in high-nutrient mesocosms had, on average, twice as many eggs as those in the low-nutrient condition, highlighting the enhanced fecundity of snails in this condition (mixed model, nutrients: F[1,56.17] � 15.88, P � 0.0001; mean eggs per egg mass � 1 SE: high nutrients, 24.8 � 1.3, low nutrients, 11.8 � 1.5; n � 115). Parasite input level did not affect snail biomass (RM-ANOVA, F[2,30] � 0.009, P � 0.60), snail egg production (F[2,30] � 0.001, P � 0.99), or chl a levels (F[2,30] � 1.772, P � 0.18).
Nutrient level and Ribeiroia egg input jointly determined the density of infected snails among treatments (Fig. 3 A and B). Increases in parasite egg input caused an increase in infected snail density; however, this effect was significantly enhanced by nutrient addition (Fig. 3 A and B). Comparable results were achieved if we used the total number of cercariae produced per sampling date or the prevalence of infection (arcsin square-root-
transformed) among snails �10 mm as response variables (see SI Fig. 5). To evaluate the effect of eutrophication on cercarial production among actively shedding snails, we conducted a mixed-model analysis with nutrient status and parasite input level as fixed factors, snail size as a covariate, and cercarial production as the response variable. Snail subjects were nested within mesocosms and within sampling date. Eutrophication significantly enhanced per capita production of cercariae (Fig. 3C), whereas parasite input and the parasite-by-nutrient inter- action had no detectable effects. Pooling across dates and parasite conditions, infected snails from high-nutrient meso- cosms produced, on average, twice as many cercariae as snails from low-nutrient mesocosms (Fig. 3C). Infected snail size was also a significant covariate in predicting cercarial production (Fig. 3C), suggesting that nutrient-mediated increases in host size facilitated increases in parasite production.
Ribeiroia egg input level and nutrient status each positively affected infection abundance in amphibians, which ranged from 0 to 48 metacercariae (n � 338) (Fig. 3D). Owing to increases in infected snail density and in per-snail release of cercariae, larval amphibians in eutrophic treatments exhibited a 2- to 5-fold increase in infection relative to amphibians in low-nutrient mesocosms. Because green frogs normally require �1 yr to metamorphose at this latitude, limb development among most animals was insuffi- cient to evaluate malformation status. Nevertheless, these infection levels are well within the range of values known to induce limb malformations in amphibians (17). Nutrient status also enhanced amphibian size (total length, ANOVA, F[1,28] � 10.728, P � 0.004), whereas parasite treatment negatively affected amphibian survival (arcsin square-root-transformed) (ANOVA, F[2,15] � 4.247, P � 0.03) (see SI Fig. 6).
Discussion Ecologists and epidemiologists are increasingly challenged to understand how large-scale agents of environmental change
Fig. 1. Experimental setup. Bird’s eye (A) and local (B) views of outdoor mesocosms used to investigate effects of nutrient enrichment on host–parasite interactions. Nutrients (N and P) and trematode eggs were added to mesocosms in a factorial experiment to understand how anthropogenic eutrophication influenced transmission of a multihost pathogen. The pathogenic trematode R. ondatrae (C, excysted metacercariae) uses pulmonate snails (D) as first intermediate hosts and amphibians as second intermediate hosts. Nutrient enrichment was hypothesized to promote algal growth, leading to an increase in the density and biomass of herbivorous snail hosts, thereby enhancing parasite transmission into snails. Infected snails with high resource availability were also expected to produce more parasite cercariae, increasing the risk of amphibian infection and pathology. In amphibians, Ribeiroia infection induces severe limb malformations (E) by disturbing the developing limb field.
15782 � www.pnas.org�cgi�doi�10.1073�pnas.0707763104 Johnson et al.
D o w
a d e d b
y g u e st
o n J
a n u a ry
1 8 , 2 0 2 2
affect host–pathogen interactions (1, 4 –7). Our study is the first to experimentally link aquatic eutrophication and transmission of a multihost parasite. In freshwater ecosystems, eutrophication is a widespread and growing problem with sharply negative effects on water quality, but the indirect effects of nutrient pollution on human and wildlife diseases are largely unexplored (8, 9, 11–14). Parasite-induced malformations in amphibians, which may have increased in prevalence and severity in recent decades (17–21), cause elevated mortality and morbidity in
affected populations, but heretofore the environmental drivers of increased infection were largely unknown (but see ref. 2). By explicitly manipulating the inputs of nutrients and parasite eggs into experimental mesocosms, our results emphasize the impor- tance of interactions among eutrophication, host dynamics, and parasite transmission. Importantly, the effects of eutrophication not only increased infection among first intermediate hosts (snails), but cascaded through the parasite’s life cycle to increase amphibian infection, thereby elevating the risk of mortality and malformation.
Eutrophication promoted parasite infection and amphibian disease risk through two related but distinct mechanisms. First, increases in primary production resulting from nutrient addition enhanced the growth, reproduction, and survival of herbivorous snails, increasing the availability of first intermediate hosts for Ribeiroia miracidia (Fig. 2). In turn, this led to an increase in the density and prevalence of infected snails (Fig. 3 A and B). Second, eutrophication caused infected snails to nearly double their individual production of cercariae relative to infected snails in low-nutrient mesocosms (Fig. 3C). This likely occurred be- cause of enhanced snail survival and growth under high-resource conditions (older and larger snails generally produce more parasites) or because of an increased capacity to translate snail resources into parasite secondary production (12, 15, 23–25). Importantly, although nutrient-mediated increases in infected host size contributed strongly to the observed increase in cer-
Fig. 2. Effects of nutrient enrichment on algal and snail growth. Nutrient enrichment significantly enhanced periphyton chl a (A) (RM-ANOVA, nutri- ents: F[1,30] � 75.93, P � 0.0001; time � nutrients: Greenhouse–Geisser cor- rected F[2.061,120] � 23.747, P � 0.0001), snail egg production (B) (RM-ANOVA, nutrient status: F[1,30] � 9.803, P � 0.004; time � nutrients: Greenhouse– Geisser adjusted F[2.58,77.39] � 6.92, P � 0.001), and dry mass of the snail host population (P. trivolvis) (C) (RM-ANOVA, nutrients: F[1,30] � 68.46, P � 0.0001; time � nutrients: Greenhouse–Geisser corrected F[2.96, 88.88] � 21.802, P � 0.0001). Snail density was converted to total dry mass by using the following equation (mass in grams � 0.0002 � [length in mm]2.7232; R2 � 0.96). Values are mean � 1 SE and are pooled among parasite egg treatments, because parasite level did not significantly affect measured response variables.
Fig. 3. Influence of eutrophication on Ribeiroia infection in snails and amphibians. Effects of Ribeiroia egg input level on the density of infected snail hosts (P. trivolvis) under high-nutrient conditions (A) and under low-nutrient conditions (B) (parasite input: F[2,30] � 18.917, P � 0.0001; nutrients: F[1,30] � 11.079, P � 0.001; parasite � nutrients: F[2,30] � 8.368, P � 0.001). (C) Influence of nutrient condition and snail size (log10-transformed) on the per capita production of cercariae by infected snails (with snails nested within mesocosm and date sampled; mixed-model analysis, nutrients: F[1,31.477] � 5.20, P � 0.03; snail size: F[1,87.09] � 28.604, P � 0.0001). (D) Mean abundance of Ribeiroia metacercariae within larval green frogs as a function of Ribeiroia egg input level and nutrient condition (ANOVA, parasite input: F[2,28] � 19.27, P � 0.0001; nutrient input: F[1,28] � 5.289, P � 0.02). No metacercariae were recovered from amphibians in the ‘‘no-parasite’’ treatment. Values are mean � 1 SE.
Johnson et al. PNAS � October 2, 2007 � vol. 104 � no. 40 � 15783
D o w
a d e d b
y g u e st
o n J
a n u a ry
1 8 , 2 0 2 2
carial production, nutrient input level also had direct positive effects on cercarial release, suggesting that our results ref lect changes in both host size and parasite productivity (26). Taken together, the increase in infected snails and in the per capita production of cercariae led to significant increases in Ribeiroia metacercariae among co-occurring amphibian larvae (Fig. 3D). Considering that most theoretical models of trematode parasites assume equivalent production of cercariae among infected hosts regardless of environmental conditions, these results have im- portant implications for understanding how environmental change indirectly affects pathogen transmission. Although the long developmental period of Rana clamitans (�1 yr) precluded inclusion of limb malformations as a response variable, the strong relationship between Ribeiroia infection and malforma- tion frequency in amphibians (2, 17–21) suggests that the observed increase in infection can reasonably be expected to increase malformation likelihood.
In nature, the relationship between eutrophication and par- asitism is likely to be variable and more complex than reported here, but several lines of evidence suggest that the proposed linkage between nutrient enrichment and Ribeiroia extends beyond the results of our experiment: (i) Available field data support correlations among eutrophication, snail host biomass, and Ribeiroia infection (19 –21). (ii) Definitive hosts (especially birds) are often attracted to eutrophic environments, thereby increasing the input of parasite eggs deposited through feces (13, 26). In our experiment, nutrient enrichment and parasite inputs were independent variables, effectively decoupling this positive feedback. If definitive hosts are also more likely to consume infected amphibians as a result of their malformations (17), this will further amplify the effect of eutrophication on Ribeiroia abundance. However, definitive host abundance and activity will also depend strongly on the availability of suitable habitat, suggesting that the effects of eutrophication on infection will be context-dependent (27). Eutrophic wetlands embedded within environments favorable to bird hosts (e.g., rangeland), for example, may be more likely to support Ribeiroia and malfor- mation epidemics than similarly eutrophic wetlands surrounded by less hospitable bird habitat (e.g., row crop agriculture). (iii) Finally, although our experiment included a simplified aquatic community, limited evidence suggests that inclusion of higher trophic levels, including snail and amphibian predators, may enhance infection through shifts in host behavior and life history strategies. By achieving size refugia, planorbid snails, such as those used by Ribeiroia, can achieve competitive dominance in high-nutrient environments (22), whereas amphibian larvae exposed to chemical cues from predators will often reduce their activity, thereby increasing their risk of trematode parasite infection (28). Nevertheless, given the inherent complexity and variability of natural communities, further study is needed to understand under what conditions eutrophication is likely to enhance infection and pathology. The current experiment in- cluded only two levels of nutrients (ambient vs. enriched), leaving open the question of whether the response of parasitism to eutrophication is linear or more complex, limiting the extent to which these results can be extrapolated over a broader range of nutrient levels.
An increase in Ribeiroia infection and malformations owing to progressive eutrophication could pose a serious risk to affected amphibian populations. Ribeiroia is a pathogenic parasite, caus- ing direct and indirect (e.g., via malformations) mortality in amphibians (e.g., ref. 16 and this study). Although our experi- ment was short-term, sustained increases in the levels of Ribei- roia infection may precipitate decline or collapse of the amphib- ian population over longer time scales. In natural environments, the overwintering of infected snails may compound the effects of eutrophication among years. Indeed, by the end of our experi- ment, the number of prepatent (immature) snails infected with
Ribeiroia more than doubled the total number of infected snails, forecasting substantially higher densities of infected snails for the following year. Because Ribeiroia causes reproductive cas- tration in infected snails (21), high levels of infection may eventually reduce the snail population, possibly leading to cy- clical patterns in Ribeiroia abundance and amphibian malfor- mations. Moreover, eutrophic habitats frequently have high levels of contaminants such as pesticides or heavy metals, which may compromise the immune resistance of amphibians and further increase their susceptibility to parasite infection (2, 29), provided that such contaminants do not adversely affect the abundance of infected snail or bird hosts. Thus, as natural wetlands continue to be altered or destroyed, amphibians, now the most imperiled class of vertebrates worldwide (30), may be increasingly forced to use marginal, often eutrophic habitats that may be hotspots for disease.
Our results have broad applicability to other multihost para- sites and their hosts. Recent increases in a variety of human and wildlife multihost parasites have been linked to eutrophication, including cholera, salmonid whirling disease, West Nile virus, coral diseases, and malaria (13, 14, 31–33). Trematode parasites similar to Ribeiroia that use snails as intermediate hosts also infect humans, ranging from the nuisance, but relatively innoc- uous, cercarial dermatitis to the pathogenic schistosomiasis, which is estimated to aff lict 200 million people across Africa and Asia (35). If the life cycles of Schistosoma spp. are similarly affected by eutrophication, forecasted increases in agricultural nutrient applications in developing countries where schistoso- miasis is endemic could hinder or inhibit efforts to control this disease. Ultimately, parasites that use multiple hosts through their life cycles are embedded in food webs with many connec- tions to environmental drivers such as climate change, nutrient mobilization, and biotic exchange (1, 5, 6, 36). Such large-scale drivers can have substantial impacts on parasites and their hosts, with consequences for plant, animal, and human health. By understanding these impacts we will be better able to forecast disease risk in a changing world.
Methods Experimental Design. To evaluate how eutrophication affected Ribeiroia transmission among hosts throughout its life cycle, we conducted a 2 � 3 factorial experiment manipulating the inputs of nutrients (ambient and elevated) and parasite eggs (none, low, and high) in mesocosms containing a community of pulmonate snails, larval amphibians, zooplankton, and algae (Fig. 1). Ex- periments were conducted in 1,200-liter mesocosms established outdoors near the University of Wisconsin Trout Lake Station (Fig. 1). Mesocosms were randomly assigned to condition and replicated six times for a total of 36 mesocosms. We initially seeded mesocosms with 1,000 liters of lake water, 22 kg of commercial ‘‘play sand’’ as substrate, and 30 g of CaCO3 to promote snail shell growth. To provide inocula of algae and zooplankton, we added 50 ml of lake sediment and 30 ml of concentrated zooplankton and phytoplankton from each of five local wetlands.
We stocked mesocosms with 50 randomly selected uninfected snails (Planorbella trivolvis) collected and pooled from three local wetlands (mean size � 1 SE � 12.8 � 0.15 mm). To avoid inadvertent introduction of infected snails into the experiment, we isolated all snails individually into 50-ml vials for 24 h and subsequently examined the associated water for trematode cer- cariae. We dissected an additional 200 snails to ensure that prepatent (immature) infections of any trematode were rare (�0.5%) and that Ribeiroia was completely absent. We collected green frog (R. clamitans) egg masses (n � 3) from three wetlands in northern Wisconsin and allowed them to hatch and develop in the laboratory [stage 26 (37)] before adding 75 randomly selected individuals to each mesocosm on July 20, 2005. Meso-
15784 � www.pnas.org�cgi�doi�10.1073�pnas.0707763104 Johnson et al.
D o w
a d e d b
y g u e st
o n J
a n u a ry
1 8 , 2 0 2 2
cosms were covered with 1-mm mesh lids to minimize coloni- zation of unintended f lora and fauna.
Experimental Manipulation: Nutrient and Parasite Additions. We experimentally enhanced N and P in half of the mesocosms by adding 4.7 g of NH4NO3 and 0.4 ml of 85% H3PO4, respectively, and left remaining mesocosms at ambient nutrient concentra- tions. These one-time additions were selected to achieve initial concentrations of �1,800 �g�liter�1 of N and 200 �g�liter�1 of P for 945 liters of water (20:1 molar ratio). One week later, after algae had begun to use nutrient resources, average total unfil- tered concentrations of N and P in the high- and low-nutrient treatments were 873 � 31.4 �g�liter�1 N and 101.8 � 3.1 �g�liter�1 P (high nutrients) and 467.3 � 2.1 �g�liter�1 N and 10.3 � 0.4 �g�liter�1 P (low nutrients), respectively. These levels are well within the range of values observed in natural amphibian habitats (19, 20). A recent survey of amphibian habitats in Wisconsin found nutrient concentrations of 2,859 �g�liter�1 N and 348 �g�liter�1 P for agricultural (eutrophic) wetlands and 1,235 �g�liter�1 N and 43 �g�liter�1 P for forested wetlands (R.B.H., unpublished data).
To obtain embr yonated eggs of Ribeiroia, we experimentally infected laborator y rats (n � 10) with 50 metacercariae isolated from infected amphibians. After 2 weeks, rat fecal matter was collected on wet paper towels, soaked in spring water for 24 h, filtered through a sieve series, and incubated in the dark at 28°C for 3 weeks (38). Water was aerated contin- uously and changed weekly. We estimated Ribeiroia egg den- sity � 1 SE by counting the numbers of embr yonated eggs in five 20-�l aliquots examined at �200 magnification. We added eggs to mesocosms biweekly at one of three levels: 0 (control), 150 � 11 (low egg input), and 1,500 � 111 (high egg input). Eggs were placed into 1-liter plastic chambers suspended at the mesocosm surface and allowed to hatch and subsequently seek out susceptible snail hosts. We used suspended chambers to ensure that eggs were exposed to sunlight, which stimulates hatching, and equipped chambers with four openings covered by a 1-mm mesh screen to allow hatching parasite miracidia to escape the chamber and enter the mesocosm. After hatching, parasite miracidia were allowed to locate and infect susceptible snail hosts, which, after a period of intrasnail maturation, developed into parasite rediae. Rediae, in turn, produced mobile cercariae that emerged nightly from infected snails and actively infected lar val amphibians, wherein they formed metacercariae. For mesocosms in the ‘‘no-parasite’’ treatment, we added a comparable volume (0.02– 0.2 ml) of sieved and incubated feces from uninfected rats to equalize any nutrient additions administered through fecal material.
Sampling Schedule and Data Collection. Biweekly throughout the summer, we quantified levels of algal growth, snail density and reproduction, snail infection prevalence, and the daily, per-snail release of cercariae. In this manner we evaluated the effects of eutrophication on Ribeiroia transmission into snails (miracidia to
rediae) and from snails into amphibians (cercariae to metacer- cariae) (see SI Fig. 4). We collected phytoplankton chl a from open water samples (2 liters) and filtered it onto 45-�m glass fiber filters, whereas we isolated periphyton chl a from 5 � 2.8-cm strips of f lagging tape suspended on the sides of each mesocosm. Filtered samples were frozen for 24 h, extracted in methanol, homogenized, and centrifuged. Chlorophyll values were measured on a spectrophotometer. Predominantly, we were interested in the effects of nutrient addition on periphyton rather than phytoplankton growth, because the former provides an important food resource for herbivorous snails. Moreover, we expected that, within our experiments, phytoplankton standing stock would be controlled by zooplankton grazing, rather than by nutrient limitation.
Over this same time period, we visually enumerated the numbers of snails, snail egg masses, and eggs per egg mass within each mesocosm. To avoid unintentional bias in data collection, mesocosms were numerically coded without any indication of treatment condition. We classified snails into four size categories (1–5 mm, 5.1–10 mm, 10.1–15 mm, and �15 mm) that could consistently be classified visually (�95% observer accuracy) and determined the number of eggs per snail egg mass by counting eggs from up to a maximum of 10 randomly selected egg masses on 11 � 17-cm Plexiglas sheets placed within each mesocosm. Beginning on July 7, 2005, we estimated snail infection preva- lence and daily cercarial production by randomly selecting 25 adult snails (�10 mm) from each mesocosm and isolating them individually into 50-ml centrifuge tubes allowed to f loat over- night within mesocosms. Released cercariae were identified and enumerated to provide estimates of snail infection prevalence and per capita production of Ribeiroia cercariae. All snails were returned to mesocosms the following morning. In total, 5,342 snails were examined for cercarial release, of which 94 were confirmed infected with Ribeiroia. Infected snails released an average � 1 SE of 83.6 � 11.2 (range: 2– 651).
Winter conditions did not allow us to maintain the experiment between years, and we destructively sampled the mesocosms on September 15. We dissected all snails �5 mm that did not release cercariae to determine the number of prepatent (immature) infections and measured (total length in millimeters) all surviv- ing amphibian larvae. We necropsied 10 larvae from each mesocosm (including individuals from the ‘‘no-parasite’’ treat- ments) to quantify Ribeiroia metacercariae.
We dedicate this research to the memory of Dr. Daniel Robert Suth- erland (1952–2006). For logistical support we thank T. Kratz, S. Knight, L. Winn, and the Wisconsin Department of Natural Resources. For assistance with experiment establishment and maintenance we thank E. Preu, A. Schiller, M. Pecore, J. Vehrs, G. Sass, J. Rusak, K. Skogen, and the Wisconsin Department of Natural Resources Division of Forestry Fire, especially the late S. Matula. K. Cottingham, K. Lunde, V. McKenzie, R. Ostfeld, V. Smith, and A. Townsend provided comments helpful in shaping the manuscript. This research was funded, in part, by grants from the National Science Foundation (DEB-0411760 and DEB- 0217533), the Anna Grant Birge Fund, and the Juday family.
1. Harvell CD, Mitchell CE, Ward JR, Altizer S, Dobson AP, Ostfeld RS, Samuel MD (2002) Science 296:2160 –2162.
2. Kiesecker JM (2002) Proc Natl Acad Sci USA 99:9900 –9904. 3. Pope K, Masuoka P, Rejmánková E, Grieco J, Johnson S, Roberts D (2005)
Ecol Appl 15:1223–1232. 4. Patz JS, Campbell-Lendrum D, Holloway T, Foley JA (2005) Nature 438:310 –
317. 5. Daszak P, Cunningham AA, Hyatt AD (2000) Science 287:443– 449. 6. Ostfeld R, Keesing F, Eviner V, eds (2007) Infectious Disease Ecology: Effects
of Ecosystems on Disease and of Disease on Ecosystems (Princeton Univ Press, Princeton).
7. National Research Council (2001) Grand Challenges in Environmental Sciences. Available at www.nap.edu/books/0309072549/html. Accessed September 1, 2006.
8. Schindler DW (2006) Limnol Oceanogr 51:356 –363. 9. Smith VH, Joye SB, Howarth RW (2006) Limnol Oceanogr 51:351–355.
10. Millennium Ecosystem Assessment (2005) Millennium Ecosystem Reports. Available at www.maweb.org. Accessed May 1, 2007.
11. Townsend AR, Howarth RW, Bazzaz FA, Booth MS, Cleveland CC, Collinge SK, Dobson AP, Epstein PR, Holland EA, Keeny DR, et al. (2003) Front Ecol Environ 1:240 –246.
12. Lafferty KD, Holt RD (2003) Ecol Lett 6:654 – 664. 13. Johnson PTJ, Carpenter SR (2007) in Infectious Disease Ecology: Effects of
Ecosystems on Disease and of Disease on Ecosystems, eds Ostfeld R, Keesing F, Eviner V (Princeton Univ Press, Princeton).
14. McKenzie VJ, Townsend AR (2007) EcoHealth, in press. 15. Smith VH, Jones TP, II, Smith MS (2005) Front Ecol Environ 3:268 –274. 16. Bennett EM, Carpenter SR, Caraco NF (2001) BioScience 51:227–234.
Johnson et al. PNAS � October 2, 2007 � vol. 104 � no. 40 � 15785
D o w
a d e d b
y g u e st
o n J
a n u a ry
1 8 , 2 0 2 2
17. Johnson PTJ, Lunde KB, Ritchie EG, Launer AE (1999) Science 284:802– 804. 18. Sessions SK, Franssen RA, Horner VL (1999) Science 284:800 – 802. 19. Johnson PTJ, Chase JM (2004) Ecol Lett 7:521–526. 20. Johnson PTJ, Lunde KB, Thurman EM, Ritchie EG, Wray SW, Sutherland
DR, Kapfer JM, Frest TJ, Bowerman J, Blaustein AR (2002) Ecol Monogr 72:151–168.
21. Johnson PTJ, Sutherland DR, Kinsella JM, Lunde KB (2004) Adv Parasitol 57:191–253.
22. Chase JM (2003) Oikos 101:187–195. 23. Pulkkinen K, Ebert D (2004) Ecology 85:823– 833. 24. Keas BE, Esch GW (1997) J Parasitol 83:96 –104. 25. Sandland GJ, Minchella DJ (2003) Oecologia 134:479 – 486. 26. Hoyer MV, Canfield DE, Jr (1994) Hydrobiologia 279-280:107–119. 27. Gagne SA, Fahrig L (2007) Landscape Ecol 22:205–215. 28. Thiemann GW, Wassersug RJ (2000) Biol J Linn Soc 71:513–528. 29. Forson DD, Storfer A (2006) Ecol Appl 16:2325–2332.
30. Stuart SN, Chanson JS, Cox NA, Young BE, Rodrigues ASL, Fischman DL, Waller RW (2004) Science 306:1783–1786.
31. Bruno JF, Petes LE, Har vell CD, Hettinger A (2003) Ecol Lett 6:1056 – 1061.
32. Cottingham KL, Chiavelli DA, Taylor RA (2003) Front Ecol Environ 1:80 – 86. 33. Reiskind MH, Walton ET, Wilson ML (2004) J Med Entomol 41:650 – 656. 34. Rejmánková E, Grieco J, Achee N, Masuoka P, Pope K, Roberts D, Higashi
RM (2006) in Disease Ecology: Community Structure and Pathogen Dynamics, eds Collinge SK, Ray CR (Oxford Univ Press, New York).
35. Steinmann P, Keiser J, Bos R, Tanner M, Utzinger J (2006) Lancet Infect Dis 6:411– 425.
36. Lafferty KD, Dobson AP, Kuris AM (2006) Proc Natl Acad Sci USA 103:11211– 11216.
37. Gosner KL (1960) Herpetelogica 16:183–190. 38. Huizinga HW (1973) Exp Parasitol 33:350 –364.
15786 � www.pnas.org�cgi�doi�10.1073�pnas.0707763104 Johnson et al.
D o w
a d e d b
y g u e st
o n J
a n u a ry
1 8 , 2 0 2 2
Are you busy and do not have time to handle your assignment? Are you scared that your paper will not make the grade? Do you have responsibilities that may hinder you from turning in your assignment on time? Are you tired and can barely handle your assignment? Are your grades inconsistent?
Whichever your reason is, it is valid! You can get professional academic help from our service at affordable rates. We have a team of professional academic writers who can handle all your assignments.
- Plagiarism free papers
- Timely delivery
- Any deadline
- Skilled, Experienced Native English Writers
- Subject-relevant academic writer
- Adherence to paper instructions
- Ability to tackle bulk assignments
- Reasonable prices
- 24/7 Customer Support
- Get superb grades consistently
Online Academic Help With Different Subjects
Students barely have time to read. We got you! Have your literature essay or book review written without having the hassle of reading the book. You can get your literature paper custom-written for you by our literature specialists.
Do you struggle with finance? No need to torture yourself if finance is not your cup of tea. You can order your finance paper from our academic writing service and get 100% original work from competent finance experts.
While psychology may be an interesting subject, you may lack sufficient time to handle your assignments. Don’t despair; by using our academic writing service, you can be assured of perfect grades. Moreover, your grades will be consistent.
Engineering is quite a demanding subject. Students face a lot of pressure and barely have enough time to do what they love to do. Our academic writing service got you covered! Our engineering specialists follow the paper instructions and ensure timely delivery of the paper.
In the nursing course, you may have difficulties with literature reviews, annotated bibliographies, critical essays, and other assignments. Our nursing assignment writers will offer you professional nursing paper help at low prices.
Truth be told, sociology papers can be quite exhausting. Our academic writing service relieves you of fatigue, pressure, and stress. You can relax and have peace of mind as our academic writers handle your sociology assignment.
We take pride in having some of the best business writers in the industry. Our business writers have a lot of experience in the field. They are reliable, and you can be assured of a high-grade paper. They are able to handle business papers of any subject, length, deadline, and difficulty!
We boast of having some of the most experienced statistics experts in the industry. Our statistics experts have diverse skills, expertise, and knowledge to handle any kind of assignment. They have access to all kinds of software to get your assignment done.
Writing a law essay may prove to be an insurmountable obstacle, especially when you need to know the peculiarities of the legislative framework. Take advantage of our top-notch law specialists and get superb grades and 100% satisfaction.
What discipline/subjects do you deal in?
We have highlighted some of the most popular subjects we handle above. Those are just a tip of the iceberg. We deal in all academic disciplines since our writers are as diverse. They have been drawn from across all disciplines, and orders are assigned to those writers believed to be the best in the field. In a nutshell, there is no task we cannot handle; all you need to do is place your order with us. As long as your instructions are clear, just trust we shall deliver irrespective of the discipline.
Are your writers competent enough to handle my paper?
Our essay writers are graduates with bachelor's, masters, Ph.D., and doctorate degrees in various subjects. The minimum requirement to be an essay writer with our essay writing service is to have a college degree. All our academic writers have a minimum of two years of academic writing. We have a stringent recruitment process to ensure that we get only the most competent essay writers in the industry. We also ensure that the writers are handsomely compensated for their value. The majority of our writers are native English speakers. As such, the fluency of language and grammar is impeccable.
What if I don’t like the paper?
There is a very low likelihood that you won’t like the paper.
- When assigning your order, we match the paper’s discipline with the writer’s field/specialization. Since all our writers are graduates, we match the paper’s subject with the field the writer studied. For instance, if it’s a nursing paper, only a nursing graduate and writer will handle it. Furthermore, all our writers have academic writing experience and top-notch research skills.
- We have a quality assurance that reviews the paper before it gets to you. As such, we ensure that you get a paper that meets the required standard and will most definitely make the grade.
In the event that you don’t like your paper:
- The writer will revise the paper up to your pleasing. You have unlimited revisions. You simply need to highlight what specifically you don’t like about the paper, and the writer will make the amendments. The paper will be revised until you are satisfied. Revisions are free of charge
- We will have a different writer write the paper from scratch.
- Last resort, if the above does not work, we will refund your money.
Will the professor find out I didn’t write the paper myself?
Not at all. All papers are written from scratch. There is no way your tutor or instructor will realize that you did not write the paper yourself. In fact, we recommend using our assignment help services for consistent results.
What if the paper is plagiarized?
We check all papers for plagiarism before we submit them. We use powerful plagiarism checking software such as SafeAssign, LopesWrite, and Turnitin. We also upload the plagiarism report so that you can review it. We understand that plagiarism is academic suicide. We would not take the risk of submitting plagiarized work and jeopardize your academic journey. Furthermore, we do not sell or use prewritten papers, and each paper is written from scratch.
When will I get my paper?
You determine when you get the paper by setting the deadline when placing the order. All papers are delivered within the deadline. We are well aware that we operate in a time-sensitive industry. As such, we have laid out strategies to ensure that the client receives the paper on time and they never miss the deadline. We understand that papers that are submitted late have some points deducted. We do not want you to miss any points due to late submission. We work on beating deadlines by huge margins in order to ensure that you have ample time to review the paper before you submit it.
Will anyone find out that I used your services?
We have a privacy and confidentiality policy that guides our work. We NEVER share any customer information with third parties. Noone will ever know that you used our assignment help services. It’s only between you and us. We are bound by our policies to protect the customer’s identity and information. All your information, such as your names, phone number, email, order information, and so on, are protected. We have robust security systems that ensure that your data is protected. Hacking our systems is close to impossible, and it has never happened.
How our Assignment Help Service Works
You fill all the paper instructions in the order form. Make sure you include all the helpful materials so that our academic writers can deliver the perfect paper. It will also help to eliminate unnecessary revisions.
2. Pay for the order
Proceed to pay for the paper so that it can be assigned to one of our expert academic writers. The paper subject is matched with the writer’s area of specialization.
3. Track the progress
You communicate with the writer and know about the progress of the paper. The client can ask the writer for drafts of the paper. The client can upload extra material and include additional instructions from the lecturer. Receive a paper.
4. Download the paper
The paper is sent to your email and uploaded to your personal account. You also get a plagiarism report attached to your paper.
PLACE THIS ORDER OR A SIMILAR ORDER WITH US TODAY AND GET A PERFECT SCORE!!!