Dietary protein restriction impairs growth, immunity, and disease resistance in southern leopard frog tadpoles
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- Venesky, M.D., Wilcoxen, T.E., Rensel, M.A. et al. Oecologia (2012) 169: 23. doi:10.1007/s00442-011-2171-1
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The immune system is a necessary, but potentially costly, defense against infectious diseases. When nutrition is limited, immune activity may consume a significant amount of an organism’s energy budget. Levels of dietary protein affect immune system function; high levels can enhance disease resistance. We exposed southern leopard frog [Lithobates sphenocephalus (=Rana sphenocephala)] tadpoles to high and low protein diets crossed with the presence or absence of the pathogenic amphibian chytrid fungus (Batrachochytrium dendrobatidis; Bd) and quantified: (1) tadpole resistance to Bd; (2) tadpole skin-swelling in response to phytohaemagglutinin (PHA) injection (a measure of the T cell-mediated response of the immune system); (3) bacterial killing ability (BKA) of tadpole blood (a measure of the complement-mediated cytotoxicity of the innate immune system); and (4) tadpole growth and development. Tadpoles raised on a low-protein diet were smaller and less developed than tadpoles on a high-protein diet. When controlled for developmental stage, tadpoles raised on a low-protein diet had reduced PHA and BKA responses relative to tadpoles on a high-protein diet, but these immune responses were independent of Bd exposure. High dietary protein significantly increased resistance to Bd. Our results support the general hypothesis that host condition can strongly affect disease resistance; in particular, fluctuations in dietary protein availability may change how diseases affect populations in the field.
Animals use a complex suite of behavioral and immunological responses to parasite and pathogen (hereafter “parasite”) exposure, ranging from parasite avoidance (Behringer et al. 2006; Rohr et al. 2009) to the production of lymphocyte and antibody-mediated defenses (reviewed in Janeway and Medzhitov 2002). Although the immune system is a critical line of defense against invading parasites, significant costs are associated with its maintenance and operation. For example, it is generally thought that development of immune responses can use a significant amount of an organism’s energy budget (reviewed in Schmid-Hempel 2003). Thus, hosts that are in poor condition (e.g., poor nutritional state) may suffer a reduced potential to mount an effective immune response. For example, independent of gonadal regression, seasonally breeding rodents that are food-deprived and losing body fat generally exhibit reduced immunity (reviewed in Demas 2004). While decreased food consumption can compromise immune function, immune maintenance and operation are strongly associated with dietary protein content in caterpillars (Spodopteralittoralis; Lee et al. 2006). Not only does high protein enhance parasite resistance (Lee et al. 2006), recent evidence shows that hosts can optimize their dietary protein intake. When challenged with a parasite infection, African armyworms (Spodoptera exempta) alter their feeding behavior such that they increase their protein intake (Povey et al. 2008).
Understanding the interplay between host condition and immune function is of great importance for amphibian conservation. Compared to other vertebrates, amphibian populations are declining at an alarming rate—nearly one-third of the described species are threatened with extinction (Stuart et al. 2004). Although a myriad of factors contribute to amphibian declines (reviewed in Blaustein and Johnson 2003; Collins and Storfer 2003), growing evidence links amphibian population declines and extinctions with infectious diseases (Daszak et al. 2003; Lips et al. 2005). Batrachochytrium dendrobatidis (Bd) is a widespread parasitic fungus which causes chytridiomycosis in amphibians and has been implicated in widespread population declines (Berger et al. 1998; Daszak et al. 2003; Skerratt et al. 2007). Bd is an aquatic parasite which infects the keratinized epithelial tissues of metamorphic and adult amphibians (Longcore et al. 1999). Severe Bd infections inhibit electrolyte transport in the epidermis of metamorphs and generally result in mortality from cardiac arrest (Voyles et al. 2009; reviewed in Voyles et al. 2011).
A growing body of evidence points towards the role of amphibian immune defenses in preventing and mitigating Bd infections (reviewed in Richmond et al. 2009; Ramsey et al. 2010), which could help explain species-specific variation in resistance. Amphibians have well-developed innate and adaptive immune responses, with the skin acting as the primary defense from infection with Bd (reviewed in Carey et al. 1999). Cutaneous antimicrobial peptides from some anuran families inhibit the growth of Bd in vitro (reviewed in Rollins-Smith and Conlon 2005; Rollins-Smith 2009), supporting the hypothesis that innate immune responses are important defenses against Bd. Less is known about specific adaptive immune responses against Bd (Richmond et al. 2009; Ramsey et al. 2010). Woodhams et al. (2007) found changes in circulating leukocytes following Bd infection of red-eyed tree frog adults (Litoria chloris). Additionally, Ramsey et al. (2010) documented Bd-specific innate and adaptive immune responses of adult African clawed frogs (Xenopus laevis) which may contribute to Bd resistance. Combined, these results support a role for host immune responses as key factors in determining resistance to Bd.
In our experiment, we examined the effects of dietary protein on growth, development, disease resistance, and immune responsiveness in southern leopard frog [Lithobates sphenocephalus (=Rana sphenocephala)] tadpoles. In tadpoles, Bd infections are restricted to the tissues around the keratinized labial teeth and jaw sheaths (Fellers et al. 2001) and tadpoles do not generally die when infected with Bd (Parris and Cornelius 2004; Garner et al. 2009; but see Blaustein et al. 2005). First, we raised tadpoles on low- and high-protein diets and tested if dietary protein content influenced their resistance to Bd. We then compared aspects of the innate and adaptive immune responses among parasite treatment groups (non-exposed, exposed but not infected, and infected) by conducting two standardized challenges of the non-specific immune system: (1) tadpole skin-swelling in response to phytohaemagglutinin (PHA) injection (a measure of the T cell-mediated response of the immune system), and (2) the bacterial killing ability of tadpole blood (a measure of the complement-mediated cytotoxicity of the innate immune system). We predicted that tadpoles from the low-protein treatment would have reduced immune capacities, ultimately resulting in decreased resistance to Bd. Also, if Bd is generally immunosuppressive, we expected to observe a weaker PHA response and a reduced bacterial killing ability for tadpoles exposed to, and/or infected with Bd, irrespective of diet treatment. We tested the effects of diet and parasite treatments, alone and in combination, on tadpole growth and development. We predicted that tadpoles infected with Bd would be smaller and less developed than control tadpoles.
Materials and methods
Animal collection and husbandry
Southern leopard frog (L. sphenocephalus) tadpoles used in our experiments were derived from eggs collected from natural ponds within Shelby County, TN, USA (35°08′16.52″N; 89°48′39.76″W). On 5 March 2010, we collected half the eggs from each of 12 L. sphenocephalus egg masses from 3 ponds within a 1.6-km radius. Immediately after collection, eggs were transported to the laboratory at The University of Memphis. Eggs and hatchling tadpoles were combined and then separated into four 37.85-L glass aquaria filled with approximately 18.5 L of aged tap water, which was continually aerated, until all tadpoles had reached the Gosner (1960) stage 25. We then haphazardly sampled tadpoles (n = 20) from each of the four aquaria to evenly distribute potential genetic effects on the larval traits we measured. Tadpoles were placed individually in 1.5-L plastic containers filled with 1 L of aged tap water. Throughout the experiment, tadpoles were maintained on a 12 L:12D photoperiod at 21°C (±1°C) and were fed approximately 0.20 g of either low- or high-protein diet every 3 days (see below).
Full water changes were conducted every 3 days. For each water change, we used a piece of mesh screen to remove the focal tadpole from its container and placed it in a temporary transfer container. We then emptied the water from the container and replaced the focal tadpole in its container, together with 1.0 L of aged tap water. After the inoculation of Bd (see below), we took the following precautions to avoid accidental parasite transmission between treatments. First, we always performed water changes on non-exposed tadpoles before exposed tadpoles. Second, we used different laboratory equipment (container, mesh screen) between each parasite treatment. At the end of the experiment, we thoroughly disinfected all containers by adding bleach (6% sodium hypochlorite) to yield a 10% solution, which kills Bd (Johnson and Speare 2003). Throughout the experiment, all equipment and water was disinfected in a similar fashion.
On 15 March 2010 (Day 0), tadpoles were haphazardly assigned to two groups (n = 40 per group) and were fed one of two experimental diets that differed in nutritional content. The isocaloric diets were custom ordered (Harlan Teklad, Indianapolis, Indiana USA) and were made of natural ingredients (e.g., fish and corn meal) and varied in protein and digestible carbohydrate content. The high-protein diet consisted of 47.6 protein, 22.5 carbohydrate, and 10.1 fat (% by weight). In comparison, the low-protein diet consisted of 13.8 protein, 48.6 carbohydrate, and 10.0 fat (% by weight). Other constituents of the diet were calcium (~1.7%), phosphorous (~1.2%) sodium (~0.35%), and potassium (~1.0%).
Batrachochytrium dendrobatidis inoculation
During February 2010, we isolated Bd from a recently dead L. sphenocephalus adult found in Shelby Co, TN, USA. Bd was grown in the laboratory on tryptone-gelatin hydrolysate-lactose (TGhL) agar in 9 cm Petri dishes according to standard protocol (Longcore et al. 1999). Prior to the use of Bd in the current experiment, we confirmed that the strain is reasonably pathogenic by experimentally infecting L. sphenocephalus adults (M. Venesky, unpublished data).
On Day 30 (14 April 2010), tadpoles from both diet treatments were randomly allocated into two parasite groups—a Bd exposed group and a non-exposed (control) group. For the Bd exposed group, we administered Bd by exposing the tadpoles (n = 40) to water baths containing infectious concentrations of fungal zoospores. We harvested Bd zoospores by adding 10.0 mL of sterile water to the cultures and collected the zoospores that emerged from the zoosporangia after 30 min. Tadpoles were placed individually in 100-mL waterbaths and exposed to 4,000 zoospores/mL for 72 h. Because the concentration of the Bd inoculate was lower than expected, we removed the tadpoles from the first inoculate and re-exposed them to a second inoculate of Bd containing 40,000 zoospores/mL for an additional 48 h. For the non-exposed group we exposed the remaining tadpoles (n = 40) to water baths with no Bd zoospores. Our design simulated transmission by water, one of the possible modes of Bd transmission in natural environments (Pessier et al. 1999).
Phytohaemagglutinin (PHA) challenge
On 10 May 2010 (Day 56), tadpoles had developed to Gosner stages 30–40 (33.97 ± 0.45, mean ± SE) and we measured the immune response of the tadpoles from the Bd exposed and non-exposed treatments (n = 20 per treatment) by challenging their immune system with a single phytohaemagglutinin (PHA) injection (Sigma-Aldrich, St Louis, MO, USA). The mean and range of Gosner stages of tadpoles in the Bd exposed and non-exposed treatments was 34.62 (30–41) and 33.88 (30–41), respectively, and did not differ significantly. PHA is a lectin derived from the kidney bean, Phaseolus vulgaris, which represents a non-specific challenge to the immune system. Injection with PHA induces a swelling around the injection site that is likely due to infiltration of T lymphocytes and other immune effector cells. A larger inflammatory response to PHA indicates a more robust immune response.
Prior to our PHA challenge, we made 1 L of amphibian phosphate-buffered saline (APBS) by adding the following ingredients to 1 L of sterilized (with a Whatman 25-mm GD/×0.2-μm pore filter) DI water: 6.6 g NaCL, 1.15 g Na2HPO4, 0.2 g KH2PO4. On 10 May 2010 (Day 56), we placed individual tadpoles in approximately 500 mL of MS222 (diluted to approximately 0.06 g/L) until the tadpole was anesthetized and lost righting ability. For accuracy, we made all measurements and injections under a Nikon® SMZ800 dissecting scope with ×10–60 magnification. We determined our injection location on each tadpole by measuring 20 mm from the posterior tip of the tail. At that location, we used a fine-gauged spessimeter (Mitutoyo; Precision Graphic Instruments, Spokane, WA, USA) to measure the thickness of the tail (skin and muscle) prior to the PHA injection. The average of three consecutive measurements was taken to minimize measurement error. We then injected 20 µl of a 1 mg/mL solution of PHA dissolved in APBS into the right side of each treated tadpole’s tail, at the junction of the tail and muscle using a 100-μL Hamilton glass syringe with a 30 G, 5/16′ disposable insulin needle. Contralateral injections of PHA were not possible given the size of anuran tadpoles. However, preliminary trials showed that the PHA dose we used elicited a measurable inflammatory response (0.427 ± 0.118, mean ± SE) that was significantly greater than that of tadpoles injected with APBS alone (0.080 ± 0.051, mean ± SE; P = 0.019). Data from a preliminary experiment showed that swelling response to PHA was greatest at 48 h post-injection. Accordingly, 48 h after the PHA, we anesthetized individual tadpoles and measured the skin swelling at the injection site as described previously. We considered the difference in skin swelling before and after injection as an indicator of the immune response in our analysis.
Bacterial killing assay (BKA)
We adapted the protocol described by Millet et al. (2007) to assay Escherichia coli (lyophilized pellets, ATCC #8739; Microbiologics, St. Cloud, MN, USA) killing ability in each tadpole. Because the effectiveness of this technique in comparing bacterial killing ability among treatments could be influenced by blood storage time after removal from the body of individual tadpoles (as in birds; Wilcoxen et al. 2010), we ensured that, for each treatment group, the length of time between the initiation of blood sampling and completion of the assay was the same (i.e. 45 min). To maintain this short time between sampling and assay, we performed four individual assays, each following collection of blood from all individuals in each treatment. We started with tadpoles from the Bd non-exposed treatment to minimize the chance of contaminating a non-exposed tadpole with Bd. The final group of assays was started within 4 h on the same day as the first group of assays, during which all experimental conditions remained the same to reduce the potential confound of treatment and time. In addition, we minimized this risk by using a unique set of control plates drawn from the E. coli stock solution at the start of the assay. Later analysis revealed no effect of starting time on the number of colonies grown on the control plates among the four assays (F3,4 = 4.129, P = 0.102).
On 12 May 2010 (Day 58), we used the remaining subset of the tadpoles from the Bd exposed and non-exposed treatments (n = 20 per treatment), which had developed to Gosner stages 32–41 (Bd exposed, high protein: 37.4 ± 2.92; Bd exposed, low protein: 33.6 ± 0.96; Control, high protein: 36.1 ± 2.03; Control, low protein: 32.7 ± 0.67; mean ± SD). To obtain blood samples, anesthetized tadpoles were patted on medical gauze to reduce the amount of water and mucosal secretions on the body, and the tail was cut off with scissors that had been sterilized with 75% ethanol. After removing the tail, we collected a blood sample of 3–7 μL (mean = 5.82 μL) in a microhematocrit capillary tube; the blood was immediately transferred by a pipettor with disposable tip to a small snap-top vial for each tadpole. Phosphate buffered saline (PBS) was then added to each sample to bring the total volume to 113 μL for use in an in vitro bacterial killing assay (BKA, see below). Blood samples were kept at laboratory temperature until their use in a bacterial killing assay. The scissors were sterilized with ethanol prior to each cut and a new pipette tip was used for each sample. We did not obtain sufficient blood from 3 of the 40 tadpoles.
Following collection of blood from all individuals in a treatment, we challenged the tadpole blood by adding 7 μL of E. coli stock solution, prepared by hydrating a single lyophilized pellet in 40 mL of PBS the day before the assays (per the manufacturer’s instructions). The 120 μL of suspension of bacteria and diluted blood was kept at laboratory temperature for 30 min to allow the challenge to proceed at near-body temperature for the tadpoles. Samples were vortex-mixed and two 50-μL aliquots of the mixture were spread onto separate trypticase soy agar plates and incubated overnight at 37°C. Controls consisted of 7 μL of the reconstituted bacteria culture diluted in 113 μL of PBS, and paired control plates were prepared for each of the four assays. For each assay, colonies were counted 24 h after placement of plates in the incubator. Bactericidal activity was calculated as the proportion of bacterial colonies killed in samples as compared to controls by dividing the difference in colony number on treatment plates and control plates by the total number of colonies on the control plates.
Because this immune response may be influenced by the number of enzymes present in the blood, we corrected for blood volumes that were lower than 7 μL (the maximum blood volume used in the assay). We considered the percentage of E. coli colonies killed (total colonies on the control plate minus corrected total colonies on the test plate divided by the total colonies on the control plate) as our indicator of tadpole immune response.
Life history responses and resistance to Batrachochytrium dendrobatidis
At the time of our immune challenges, we also recorded the total length (TL) (to the nearest 0.01 mm) and Gosner stage of all individuals. After completing the immune challenges, we sacrificed all tadpoles, dissected their mouthparts, and stored mouthparts in 100% EtOH for qPCR analysis to confirm tadpole infection status. DNA was extracted from mouthparts using Qiagen DNeasy kits. Quantitative PCR was done adapting methodology from Boyle et al. (2004). Primers and probe concentrations were identical, but samples were instead run using Applied Biosystems Taqman Fast Master mix in 10 µL reactions including 3 μL of DNA template. All samples were done in triplicate and were rerun if only one of the three of the wells was positive. We screened all of the tadpoles from the Bd-exposed treatment and randomly selected half of tadpoles (n = 20) from the non-exposed treatment.
We used MANOVA to test for the effects of independent variables diet (low and high protein) and parasite treatment (non-exposed, exposed but not-infected, and infected) and their interactions on the dependent variables size (TL) and stage (Gosner). We then used Bonferroni-adjusted (significance level of 0.025 for each response variable) ANOVA contrasts on each response variable to determine significant contributors to multivariate effects. Since tadpoles fed a high-protein diet were significantly more developed than tadpoles fed a low-protein diet (see “Results”), we used Gosner stage as a covariate in all of our analyses to control for potential differences in immune response as a function of developmental stage. For the PHA challenge, we used ANCOVA to test for the effects of the independent variables diet (low- and high-protein) and parasite treatment (non-exposed, exposed but non-infected, and infected) on the dependent variable skin swelling. We used log-transformations to normalize skin swelling responses. For the BKA assay, we used ANCOVA to test for the effects of the independent variables diet (low- and high-protein) and parasite treatment (non-exposed, exposed but non-infected, and infected) on the dependent variable (% of E. coli colonies killed). Within the Bd-exposed treatment, we used a 2 × 2 contingency table to test whether the proportion of tadpoles that became infected differed between the high and low protein treatments.
Effect of diet on disease resistance
There was no mortality during the experiment. No individuals from the sub-sample of tadpoles in the control treatment tested positive for Bd. Among tadpoles exposed to Bd, dietary protein content significantly affected resistance to Bd. Tadpoles raised on a low-protein diet showed a significantly higher prevalence of infection, with 65% of the tadpoles raised on a low-protein diet infected with Bd compared to 15% of tadpoles raised on a high-protein diet (χ2 = 10.42, P < 0.001).
Effects of diet and Batrachochytrium dendrobatidis on immune responses
Effects of diet on larval life history traits
Poor host immune function can lead to decreased resistance to parasitic infections (Houdijk et al. 2000; Povey et al. 2008) and higher parasite-induced mortality (Peck et al. 1992). For example, ruminant mammals are often exposed to a great diversity of parasitic trematodes and nematodes; high protein dietary supplementation often increases parasite resistance in livestock (reviewed in Coop and Kyriazakis 2001). As predicted, we found strong effects of dietary protein content on tadpole resistance to Bd. A greater proportion of tadpoles raised on the low-protein diet became infected with Bd compared to tadpoles raised on a low-protein diet (65 vs. 15%, respectively). In addition to dietary protein levels, other factors can influence host condition and ultimately lead to differences in resistance. Poor overall body condition can decrease resistance; for example, Garner et al. (2009) found that metamorphs in poor condition were less resistant to Bd-induced mortality.
Results like those of Garner et al. (2009) may occur because poor overall host condition reflects nutritional condition, and inadequate nutrients or low energy can suppress the resources devoted to immune maintenance and deployment (Lee et al. 2006). Decreased dietary protein can suppress specific immune functions (Lochmiller et al. 1993). Tadpoles have a well-developed immune system that includes lymphocyte activity (Rollins-Smith 1998); however, the effects of host nutritional status has on specific tadpole immune responses have not been explored. An effective immune response (e.g., lymphocyte recruitment and antibody production) requires an increase in metabolic costs due to increased protein synthesis (Borel et al. 1998). We found that tadpoles raised on a low-protein diet had lower PHA-induced skin-swelling and bacterial killing ability than tadpoles raised on a high-protein diet. Our results suggest that tadpoles raised on a low-protein diet were protein deficient, and were therefore unable to deploy an effective response to the PHA injection. Several components of the innate immune system may mediate the ability of tadpole blood to kill E. coli, which may also be influenced by dietary protein content. Since several responses of innate immunity (e.g., phagocytes, natural antibodies, and complement; Millet et al. 2007) may be measured in the BKA, we cannot ascertain which specific component(s) results in enhanced bacterial killing in this assay. However, our data support the findings of other studies in that organisms that have a strong bacterial killing ability can resist parasitic infections (Townsend et al. 2010) and potentially increase their survival (Wilcoxen et al. 2010). It is important to note that, by assaying all the tadpoles from the Bd non-exposed treatments first, we could have confounded treatment with time. However, we found very little inter-variation in the number of E. coli colonies that grew on the control plates as a function of assay starting time. In addition, we would have expected a directional trend in bacterial killing ability (either increased or decreased) relative to the assay starting time, which we did not find. Thus, we are confident that our data reflect treatment differences in dietary protein.
If Bd is generally immunosuppressive, we expected to observe reduced immune parameters (i.e. weaker PHA response and a reduced bacterial killing ability) for tadpoles exposed to and/or infected with Bd. Ongoing studies in the Rollins-Smith laboratory suggest that factors produced by Bd can inhibit lymphocyte proliferation (J. Ramsey, unpublished data). If those factors were taken up by the tadpole circulatory system, we might have expected to observe suppression of PHA-induced tail swelling or in the bacterial killing assay. However, we did not measure any significant effect of Bd on immune responses.
Recent studies show that amphibians use components of their innate and adaptive immune system in response to Bd infection (Rollins-Smith and Conlon 2005; Woodhams et al. 2007; Ramsey et al. 2010). For example, Ramsey et al. (2010) found that adult African clawed frogs) have Bd-specific antibodies (IgM and IgY) and antimicrobial skin peptides, both of which are effective immune defenses against Bd. However, these studies examined the immune defense against Bd infections in adult amphibian epidermis. Immune defenses around the oral apparatus of tadpoles are not well studied, but cell-mediated immunity and/or mucosal antibodies could play a role in clearance of Bd-infections in the mouth. Given the effect of high-protein on increased Bd resistance, it is likely that some component of tadpole immune system aided in Bd resistance.
It is important to note that, because we assessed infection status at the end of the experiment, we are unsure if Bd-negative tadpoles were not infected with Bd because they resisted Bd or cleared their infection during the duration of the experiment. It is possible that the immune defenses of tadpoles may have prevented Bd from infecting their keratinized mouthparts. Tadpoles might also have mounted an immune response against Bd after they became infected, clearing the infection. Regardless, our data clearly show that dietary protein content can influence the outcome of parasite exposure and are consistent with the hypothesis that there are high constitutive costs of maintaining immune function (e.g., Alaux et al. 2010).
Although we did not measure other aspects of tadpole physiology, it is possible that poor nutrition elevated corticosterone (the primary anuran stress hormone) levels, which may have inhibited immune responses in tadpoles fed a low-protein diet. For example, when nutritionally restricted, couch’s spadefoot toad (Scaphiopus couchii) tadpoles exhibited higher levels of whole-body corticosterone (Crespi and Denver 2005; Ledon-Rettig et al. 2009). Because the hypothalamic–pituitary–interrenal (similar to the hypothalamic–pituitary–adrenal axis in mammals) and immune system function are coupled (Rollins-Smith 2001), it would not be surprising if increased corticosterone mediated either of the immune responses that we measured. Specifically, increased corticosterone reduces circulating lymphocytes in Xenopus tadpoles undergoing metamorphosis (Rollins-Smith et al. 1997), which could ultimately inhibit PHA-induced skin swelling.
Given that tadpoles infected with Bd alter their feeding kinematics (Venesky et al. 2010) and obtain less food (Venesky et al. 2009) compared to non-infected tadpoles, we did not find an effect of Bd infection on tadpole growth and development. We suspect that a combination of the Bd dose and the duration of the experiment were not sufficient to reduce growth and development in this species. For example, Rachowicz and Vredenburg (2004) found that Lithobates spp. infected with Bd begin to lose keratin 49 days post-infection, whereas Bd-induced damage to Bufo and Hyla spp. can occur within ~21 days (M. Venesky, unpublished data). Indeed, the tadpoles in this study had a low incidence of mouthpart deformations. Thus, it is likely that the Bd infections were not severe enough to inhibit normal feeding and did not significantly reduce their growth and developmental.
In addition to the effects of nutrition on tadpole immune function and resistance to Bd, our results provide key insights into how specific components of tadpole diet contribute to their growth and development. We found that tadpoles raised on the high-protein diet were larger and more developed than tadpoles raised on the low-protein diet. While detailed information on diets is lacking, tadpoles have access to a variety of food items in the aquatic environment that likely differ in nutritional quality. For example, nitrogen-fixing blue-green algae (Pryor 2003) and animal products (Altig et al. 2007), each of which are high in dietary protein, can make up a significant amount of a tadpole’s diet. Thus, species differences in immune responses of tadpoles and their resistance to infectious diseases may be driven, in part, by the biotic make-up of their aquatic environments.
Resisting parasitic infection can use a substantial amount of an organism’s energy budget. If certain macronutrients, such as protein, are missing from an organism’s diet, they may suffer a reduced potential to mount an effective immune response. In our study, we found that reduced dietary protein content can impair host immune function and increase disease risk. Our results underscore the importance of host condition on disease resistance and suggest that variation in dietary protein availability may, in part, influence patterns of disease prevalence in the field.
We thank F. Brem for assistance collecting anuran eggs. The Rohr Lab at The University of South Florida provided helpful comments on an earlier draft of this manuscript. We also thank R. Alford for providing extensive editorial comments on an earlier draft of this manuscript. Collection permits from Tennessee were obtained prior to collecting the animals used in these experiments and all experimental procedures were approved by the University of Memphis IACUC. The experiments comply with the current laws of the USA. This publication was developed, in part, under a GRO Research Assistance Agreement No. MA-916980 awarded by the U.S. Environmental Protection Agency to M. Venesky. It has not been formally reviewed by the EPA. The views expressed in this document are solely those of the authors and the EPA does not endorse any products or commercial services mentioned in this publication. T. Wilcoxen was supported by a NSF Doctoral Dissertation Improvement Grant (IOS-0909620). L. Rollins-Smith was supported by NSF grants IOS-0619536 and IOS-0843207. Bd qPCR analysis was performed on instrumentation provided by NSF (MRI-0923419) to J. Kerby.