Physiology of Herpetofauna in an Urban Ecosystem

Purpose

The propose of this project is to compare amphibian and reptile assemblages in the city parks of and rural localities across Pennsylvania. The main project goal is to investigate the physiology of herpetofaunal species across a gradient of urbanization in Pennsylvania. Comparing these sites will give an idea of how urbanization has altered the diversity and physiology of herpetofauna as a result of living in an urban ecosystem.

Background

Roughly half of the world’s population lives in urban areas, but this is expected to increase to more than two-thirds by 2050 (United Nations, 2019). The expansion of urban habitats has had an overall homogeneous impact on biodiversity by filtering out species that cannot survive urban life (McKinney, 2002, 2006). An important mechanism that affects wildlife in urban areas is climate alterations that cause higher temperatures for longer periods compared to rural areas. This is called the ‘Urban Heat Island Effect’. (Imhoff et al., 2010). However, it is unclear how species that survive in metropolitan areas will cope with the potential adverse effects of urban heat islands. Therefore, the urban heat island offers an opportunity to study how and by what mechanisms organisms have responded to recent man-made changes to the environment. Physical changes associated with urbanization have been observed in several taxa (Johnson and Munshi-South, 2017), but little is known about the degree these physiological traits change in urban animals, especially amphibians and reptile, which are heavily affected by temperature changes. I will compare the physiology (field body temperatures, metabolic rates, heat tolerances, and gut microbiota) in urban and rural populations of Pennsylvania herpetofauna. I predict that organisms in the urban localities will have adaptive responses to urbanization that are similar throughout the herpetofauna community (Brans et al., 2017), with possible outcomes, including lower metabolic rates at higher temperatures, more variable body temperatures, greater temperature tolerances, and greater homogeneity of microbiome diversity in urbanized locations.

I will study herpetofauna across an urban-rural gradient in Pennsylvania, with three urban centers that are roughly equal in latitude but vary in size of human population: Philadelphia (1.6 million people), Pittsburgh (300,000 people) and Harrisburg (50,000 people). These urban locations are paired with rural areas of similar latitude which will most likely have the same species: Powdermill Nature Reserve, Letterkenny Army Depot, and Crow’s Nest Preserve. The herpetofauna will be captured by hand through visual-encounter surveys during five day sampling windows from April to October. I will capture up to 10 animals from each species we encounter in each unique location. Based on civilian scientific data on the most common herpetofauna species in these urban areas (inaturalist.org), we expect to encounter up to three species of salamanders (Plethodon cinereus, Desmognathus fuscus, and Desmognathus orchophaeus), four species of snake (Thamnophis sirtalis, Nerodia sipedon, and Storeria dekayi, and Pantherophis spiloides), and three frog species (Anaxyrus americanus, Lithobates catesbeianus, and Lithobates clamitans). Surveys entail visual inspection of the habitat and flipping of logs, rocks, and other objects to find and collect animals. The study will cover the habitats where the species are most likely to occur, as well as at the appropriate time of day and season when the species is most likely to be active. This project will survey for 10 of the most common herpetofauna species in Pennsylvania which are widespread in the northeastern United States and have large and stable populations, and therefore all of them are currently registered as species of Least Concern by the IUCN Red List of Threatened Species (iucnredlist.org).

Animals found in the studies are captured, body temperature is measured (Huey, 1974; Huey and Webster, 1976), and they will be placed in a sterile container in the field until a fecal sample can be obtained. The sample will be collected, stabilized with ethanol and placed on ice until it can be taken to a lab to analyze its microbiome.

Once fecal samples are obtained, the animals will be taken to a laboratory and used to test for temperature tolerances or to measure metabolic rate. Note that no animal will undergo both these protocols as one test may affect the results of the other. For temperature tolerance experiments, the animals acclimatize in the laboratory to room temperature of 20 °C for a specified time e.g. 24 hrs. (Alexander et al., 1999). The animals are then tested with a non-lethal dynamic temperature rise protocol to assess the critical thermal maximum (CTmax) and critical thermal minimum (CTmin), two physiologically significant tolerance limits. (Lutterschmidt and Hutchison, 1997; Angilletta et al., 2007; Camacho et al., 2017). CTmax and CTmin are measured when the animals lose their righting response, which is defined as the time when it cannot return to its feet within five seconds after being placed upside down. (Christian et al., 1988; Huang et al., 2007; Catenazzi et al., 2014; Diamond et al., 2017, 2018). Each individual is placed in a thin plastic container with a layer of water 3–5 mm deep, the containers are then placed in a water bath (Navas, 1997; von May et al., 2017). In the case of CTmax, the bath temperature will gradually rises from 20 ° C to 40 ° C at a rate of 1 ° C / min with the addition of hot water. The temperature of the CTmin will be gradually lowered from 20 ° C to 0 ° C with the addition of ice to the water bath. I will place the animals in an upside down position and when they can now longer right themselves within five seconds, I will record their temperature. I will measure the animal’s body temperature using a thermometer placed into the cloaca. The self-righting response is important when considering selection on thermal physiology because an animal that is unable to display their self-righting reflex will likely be unable to escape predation. In addition, temperature tolerance measurements have been shown to produce reproducible measurements in physiological experiments (Morgan et al., 2018).

For metabolic rate measurements, animals will fast for a specified amount of time, likely 3 to 6 days depending on body size, at a room temperature of 20 °C before the testing (Gifford, 2016). Fasting is necessary because for metabolic rate measurements, individuals need to be post-absorptive during testing (Dorcas et al., 2004). They will be tested for standard metabolic rate (SMR) by placing them in sealed off respirometers or metabolic chamber to measure oxygen consumption rates using a free flow system (Homyack et al., 2010). Animals will be given an acclimation period to adjust to the metabolic chambers and room temperature prior to acquiring measurement data. Animals will be exposed to 5°C temperature levels from 5 and 35 °C (McCue and Lillywhite, 2002). The experimentation order for these temperatures will be randomized to control for any potential effects of temperature on acclimation rates. Oxygen consumption rate will be taken every 24 minutes for 24 hours (Toledo et al., 2003; Zaidan, 2003).

After testing, the animals will be humanely euthanized and prepared as specimens using the American Society of Ichthyologists and Herpetologists guidelines for collection of live amphibians and reptiles in field and laboratory research (Beaupre et al., 2004). Animals to be prepared as specimens will be euthanized using guidelines from the AVMA Guidelines on Euthanasia (AVMA, 2013), followed by fixation in a 10% formaldehyde water solution. Liver or muscle tissue samples will be taken for DNA analysis and then placed in small glass vials containing concentrated ethanol. After all research have been conducted, specimens and tissue samples will be donated to a Pennsylvania museum or university for long-term storage.

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