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Recent papers: Red eared terrapins and alpine newts in Scotland and their possible impact on common frog tadpoles

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Written by Dr Andrew Smart, Conservation & Science Manager

A paper presented by Chris Catherine at the 2023 Glasgow Natural History Society Scottish Herpetological Conference has recently been published online in the Glasgow Naturalist. The paper outlines information about Scottish invasive species from a range of data sources. Catherine highlights the confusion around the status of the barred grass snake (Natrix helvetica) in Scotland with records from Dumfries and Galloway possibly of native animals or from a feral population. Catherine found 101 records of four different non-native species: alpine newt, sand lizard, Chinese pond turtle and common slider. The only record of sand lizards in Scotland is a single record form1970 on the island of Coll as part of a benign introduction programme in the 1970s, to help protect the species in the UK.  Similarly, Chinese pond turtle has only been recorded once, at Glasgow botanic gardens in 2022.

The alpine newt is a concern because of its ability to act as a vector for Batrachochytrium fungi the cause of chytridiomycosis or chytrid fungus in our native amphibians. In Scotland there appear to be four distinct populations. 

Common slider, the red-eared terrapin, is the most widely distributed invasive herpetofauna species in Scotland with records from 42 10km grid squares and 10 distinct populations. Globally, where the species reproduces it can cause significant impacts and is listed by the IUNC as one of the 100 most invasive species.  There is no breeding recorded in Scotland but  these animals can live for 20-40 years

Common slider/ red-eared terrapin, is the most widely distributed invasive herpetofauna species in Scotland

A second recent paper by Vodrazkova et al, working in the Czech republic undertook a lab based study on the impact of red eared sliders on frog tadpoles. Following up on previous research, Vodrazkova et al examined the speed of metamorphosis, size of froglets and the level of stress hormone in frog tadpole in a lab environment with or without the presence of  a red eared terrapin in the water (isolated form tadpoles but present in the tank).

The result showed that long-term and short-term presence of the terrapin in the same water increased the stress levels within the tadpoles, with hormone levels indicating the presence of chronic stress when the terrapin was present.  The tadpoles in the three treatments showed no significant difference in time taken to metamorphose but the size of the froglets in the three treatments varied, with smaller animals at metamorphosis in the treatments in the presence of the terrapin.  This indicates a developmental strategy to metamorphose at the smallest possible size to avoid the risk of predation in the waterbody.

At the moment red eared terrapins are not breeding in the UK but with a significant range, there is a risk that temperature increase due to global warming could lead to a new more widespread threat to our native species in the future.

Bare hands, gloves or not at all – what’s best for amphibians and why?

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Written by Andrew Smart, Conservation and Science Manager

It’s not easy working with amphibians and controlling the desire to pick them up and look at them. …from our childhood, the excitement of catching a frog, rescuing a toad or catching a newt in a pond are feelings we carry with us into adulthood. Amphibians are wonderful animals, we want to look at them, observe them and hold them in our hands.    But what happens when we do that?

What does it mean for the animals and why do we need to pause and remind ourselves the best thing to do is not handle amphibians at all?

The Amphibian and Reptile groups of Great Britain advice note 4 (ARG, 2017) suggests that amphibian workers should:

  • Handle [animals] with powder free disposable vinyl gloves which are rinsed in pond water before contact with animals.
  • A single set of gloves can be used when working at any site but should be disposed of between sites. 
  • Equipment, including footwear, nets, traps etc should also be disinfected between sites.

 

The Herpetofauna Workers Manual (revised 2003) : suggests that:

  • handling is kept to an absolute minimum because of the impact of the warmth of the human hand on the animal
  • if necessary, “good practice [is] to wet hands with pond water before handling amphibians”. 

The risk of transmission of disease such as ranavirus or chytridiomycosis (chytrid fungus) caused by Batrachochytrium dendrobatidis (Bd) and Batrachochytrium salamandrivorans (Bsal) between sites remains high as spores can survive on equipment, footwear or vegetation. Chytrid causes death by disrupting the uptake of salts through the skin, disrupting amphibian metabolism and ultimately stopping their hearts from beating (ARC, 2024). The ‘perceived wisdom’ is that handling with dry hands can damage the mucus layer, hence the need to wet either hands or gloves prior to handling an amphibian.

Varga et al (2019) highlighted the value of amphibian skin as an ‘innate immune organ’ acting as the first line of defence against pathogens in an aquatic environment.  Amphibians have a glandular network beneath their skin that produces antimicrobial and toxic chemicals which help protect the animal against pathogens and predators. Mucosal glands produce a mucus layer which protects the skins, maintaining elasticity, permeability and moisture on the skin surface (Varag et al, 2010). Along with acting as a physical barrier between amphibians and their environment, the mucus layer is a matrix containing bacteria and antimicrobial, antibacterial, antiviral, antifungal or anti-parasite secretions from the animal itself. In many species the mucus also contains alkaloids, secreted as a defence against predation. Damage to this mucus layer can have an impact on animals; handling can remove the mucus layer and some chemical pollutants, such as pesticides, can damage skin mucus having a sub-lethal effect on amphibians in both larval and adult forms.

A recent review by Woodhams et al (2023) considered the ‘adaptive microbiome hypothesis’ arguing that amphibians, like other vertebrates (including humans) have large numbers of bacteria on their skin. Previously, Rollins-Smith (2009) found beneficial bacteria within the skin mucus of amphibians produce antifungal compounds which inhibit the growth of Bd and suggested that bio-augmentation could be a future possibility to protect highly endangered species.  Woodham’s (2023) review found evidence that while Bd was increasingly virulent as it passed through Central America; in Panama, amphibian ‘mucosal skin defences’ were higher after the disease appeared, indicating a possible response by amphibian populations as their microbial community evolved to combat Bd.  The hypothesis proposed suggests that on recovery from an initial infection, the microbial community can better combat subsequent exposures.  This could also explain why some amphibians can operate as carriers of Bd and Bsal fungus while others suffer significant mortality on exposure.

In 2010 Phillott et al highlighted the need to minimise exposure of amphibians to pathogens during any form of field study, emphasising the need for strict bio-security and hygiene when moving between sites, including:

  • disinfecting footwear using sodium hypochlorite
  • disinfecting equipment such as scales, callipers or containers

Phillott et al (2010) also discussed the damage that can result from handling amphibians and identified a number of papers that found stressed animals had a greater risk of infection.  Apart from the impact on their skin, handling amphibians can cause the stress, and repeated handling has been shown to impact their behaviour.  Bliley and Woodley (2012) studied the impact of repeated handling in a species of salamander and found that repeated handling reduced feeding behaviour in females and activity in both males and females. They argued that stressed animals may even avoid reproduction and divert energy into immediate survival.

Phillott et al (2010) argued the direct transfer of microbes and spores between animals can be reduced by using single-use gloves. Importantly, they highlighted previous work that found some tadpoles suffer lethal and sublethal effects when exposed to latex (Gutleb at al 2001) and nitrile and to a lesser extent vinyl so larvae should only be handled with gloves that have been proven safe for that species or with hands wetted by water from the source pond. They argued that when gloves are not available, hand washing with 70% ethanol allowing the hands to air dry between handling is an alternative. If this is not possible in the field, ‘hand washing in the water body to which the amphibian is naturally exposed’ is the best alternative (Phillott 2010).

There seems to be some variation in the impact of gloves; Mendez et al (2008) found that water washed from nitryl gloves killed Bd spores on contact while latex and polyethylene gloves had no effect. It is important to note the effect of vinyl gloves varied with brands and batches. Human skin also demonstrated a fungicidal effect on Bd, possibly due to the presence of antifungal peptides on the skin surface.  They recommended that to minimise the risk of transfer of disease between individuals single use disposable gloves were the best option followed by bare hands as a preferable alternative to continual use of the same pair of gloves (Mendez 2008).

A more recent study (Thomas et al, 2020) in evaluating the impact of nitrile and vinyl gloves on Bd and Bsal found glove ‘rinsewater’ led to a reduction in Bd and Bsal spore production compared to water alone. They recommended:

  • the single use of powder-free vinyl gloves should be advocated to handle amphibians.
  • at a minimum gloves should be changed between each population
  • the repeated use of the same pair of gloves should be preferred above handling animals bare handed

So we know the value of the mucus layer on the skin of an amphibians and we understand the risk of damaging that skin, disrupting the animal’s immune defences.  We are also aware of the risk of disease transmission from animal to animal if we handle multiple animals in the field. There remains a dilemma as to which advice to follow; the 2017 guidance aligns with the most recent experimental work but how does a toad patroller change their gloves between each animal and still manage to collect the animals before they reach the road verge?

We have to agree a sensible approach; animals in the same population share the risk of disease transfer within their aquatic environment and so the existing 2017 ARG Advice Note 4 appears to remain the most sensible approach to handling animals. However, on some occasions, if necessary, a hand dipped in the source water-body may do less damage than not moving the animal. 

Ideally, we should do our best to restrain our excitement and look rather than touch!

 

References:

ARC [online] accessed 2024 Chytrid fungus | Amphibian and Reptile Conservation (arc-trust.org)

ARG (2017) [online] accessed 2024 4. Advice note 4 (revised) – Amphibian Disease Precautions, A Guide for UK Fieldworkers.pdf – Amphibian and Reptile Groups of the UK (arguk.org)

Bliley, J.M. and Woodley, S.K., 2012. The effects of repeated handling and corticosterone treatment on behavior in an amphibian (Ocoee salamander: Desmognathus ocoee). Physiology & behavior, 105(5), pp.1132-1139.

Gent, T. and Gibson, S. eds., 2003. Herpetofauna workers’ manual (rveisd reptint) . Peterborough: Joint Nature Conservation Committee. Herpetofauna Workers’ Manual (revised reprint) | JNCC Resource Hub

Gutleb, A.C., Bronkhorst, M., van den Berg, J.H. and Murk, A.J., 2001. Latex laboratory-gloves: an unexpected pitfall in amphibian toxicity assays with tadpoles. Environmental toxicology and pharmacology, 10(3), pp.119-121.

Mendez, D., Webb, R., Berger, L. and Speare, R., 2008. Survival of the amphibian chytrid fungus Batrachochytrium dendrobatidis on bare hands and gloves: hygiene implications for amphibian handling. Diseases of aquatic organisms, 82(2), pp.97-104.

Phillott, A.D., Speare, R., Hines, H.B., Skerratt, L.F., Meyer, E., McDonald, K.R., Cashins, S.D., Mendez, D. and Berger, L., 2010. Minimising exposure of amphibians to pathogens during field studies. Diseases of aquatic organisms, 92(2-3), pp.175-185.

Rollins-Smith, L.A., 2009. The role of amphibian antimicrobial peptides in protection of amphibians from pathogens linked to global amphibian declines. Biochimica et Biophysica Acta (BBA)-Biomembranes, 1788(8), pp.1593-1599.

Thomas, V., Van Rooij, P., Meerpoel, C., Stegen, G., Wauters, J., Vanhaecke, L., Martel, A. and Pasmans, F., 2020. Instant killing of pathogenic chytrid fungi by disposable nitrile gloves prevents disease transmission between amphibians. Plos one, 15(10), p.e0241048.

Varga, J.F., Bui-Marinos, M.P. and Katzenback, B.A., 2019. Frog skin innate immune defences: sensing and surviving pathogens. Frontiers in immunology, 9, p.433806.

Woodhams, D.C., McCartney, J., Walke, J.B. and Whetstone, R., 2023. The adaptive microbiome hypothesis and immune interactions in amphibian mucus. Developmental & Comparative Immunology, 145, p.104690.

 

Croaking Science: Amphibians and Chemical Pollution

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Written by Dr Andrew Smart, Conservation and Science Manager

Concern over the impact of chemicals on the environment began in the 1960s following the publication of ‘Silent Spring’ by Rachel Carson, which highlighted the risks of using pesticides such as DDT. Support for agriculture through the ongoing use of pesticides, herbicides and chemical fertilisers continues to raise concerns because of the impacts on beneficial invertebrates and lower vertebrates such as amphibians and fish, leading to ‘build-up’ within food chains.

Chemical impacts on amphibians result from terrestrial spraying and from the acute and chronic effects of chemicals being carried into ponds and wetlands. Amphibians have life cycles that involve changes of form when in an aquatic environment: from egg to larvae and larvae to adult, and then a terrestrial phase as adult, returning to wetlands to breed. Chemical pollutants that have been found to impact amphibians include pesticides, herbicides, fertilisers, de-icing salts and road materials, heavy metals and lead, and further impacts from microplastics have been identified. Little work has been undertaken on UK species, with the bulk of ecotoxicological work undertaken in the USA and Europe, on species often similar to and sometimes of the same genus as UK species of newts, frogs and toads.

A review of the impact of agricultural chemicals on amphibians in 2009[1] found that interactions between agrochemicals and other environmental factors impacted amphibians, disrupting thyroid activity and other biochemical pathways, impacting metamorphosis and sexual differentiation. Exposure to insecticides led to impaired behavioural responses, impaired growth, delayed metamorphosis and increased use of detoxification pathways which could impact the immune repose to other factors because of increased energy use. These ‘sub-lethal’ effects are the focus of much of the recent research.

In 2012[2] and 2013[3] analyses of previously published work found the overall effects of pollutant exposure included a reduction in amphibian survival body mass and an increase in the frequency of abnormalities during development and metamorphosis. The measured impacts were found to lead to as much as 14% reduction in survival, 7.5% decrease in body mass and a 5 fold increase in abnormalities. Pollution appeared to have no impact on the time from spawning to hatching or the time to metamorphosis but were again found to lead to altered behaviour, changes to metabolism, reproductive failure and increased embryonic and larval mortality. Immunotoxicity was found to generate a sub-lethal effect leading to higher parasite load and subsequent population decline. Road de-icers were more toxic than nitrogenous compounds while pesticides and wastewater pollution caused an intermediate reduction in survival.

A further meta-analysis[4] in 2013 found that pesticides and fertilisers had a significant negative effect on survival and growth across all the amphibian species studied with marked differences between chemical classes in their impact on amphibians: many inorganic fertilisers, insecticides and herbicides negatively affected amphibian survival, while some negatively affected larval growth.

 

Pesticides

Investigation in the USA[5] found complex mixtures of pesticides from wetlands including up to 8 fungicides, some previously unreported in tissue. Many studies describe wastewater contaminants as a significant threat to amphibian species based on their lethal and sub-lethal effects; amphibians can be subject to damage through skin and food ingestion[6]. Agricultural chemicals used as individual ingredients and mixed together led to sub-lethal changes in stress levels and physiological responses in amphibians[7].

A 2023 meta-analysis investigating the impact of different types of disturbance on reptiles and amphibians found the greatest impact occurred from drought, followed by invasive species and then agriculture. The review[8] found that agriculture had a moderate to large negative impact on body condition linked to low quality habitat, lack of foraging habitat and shelter on the impact of chemical contaminants. Many studies were listed indicating negative effects of pesticide exposure on embryos and developing tadpoles, including deformities, arrested development, mortality, endocrine and immune disruption, and increased disease susceptibility. Pesticide residues are broken down through processes into various ‘transformation products’[9] which have been detected in groundwater and surface water. Investigation of the effects of widely used pesticides and their transformation products in cell cultures found that approximately 50% of the transformation products exhibited stronger effects than the original pesticides.

Endosulfan, an organochlorine pesticide that is found in many ecosystems, including wetlands particularly in proximity to agriculture[10] , has been found to have a sub-lethal effect affecting tadpole activity and survival. In amphibians, pesticide exposures have been shown to alter biochemical processes [11], reduce tadpole survival to < 1%, survival to 10%[12] and impairment of mobility 12. Another pesticide, chlorpyrifos, has also been shown to alter tadpole brain development[13] .

Other pesticide compounds, also found in products including detergents, plastics, emulsifiers and cleaning products, impact newt physiology, disrupting reproduction, metamorphosis and the ability of amphibians to respond to changing environments[14] and  may also induce organ toxicity[15]. Pesticide linked compounds that increase the ability to ‘stick’ to surfaces, at sub-lethal concentrations have also been found to create physiological changes in amphibians[16] and some affect hormone systems with feminising effects on male gonad development identified in one study in the USA. [17] 

Pesticides can have other effects with disruption of pheromone communication in newts leading to a delayed response to female odours and, consequently, a reduction in mating success[18]. The impact is not restricted to larvae or aquatic systems, a study on the effects of seven pesticide products on juvenile European common frogs (Rana temporaria) in an agricultural overspray scenario found mortality ranged from 100% after one hour to 40% after seven days at the recommended label rate of currently registered products[19].

 

Non-agricultural pesticides

In the UK, chemicals that are banned for routine agricultural purposes are sold as parasiticide treatments for pets[20] and can enter the natural environment. Their active ingredients are toxic to many freshwater species and concentrations found from field studies are likely to negatively affect aquatic life, impacting food webs and amphibians in urban and semi-urban ponds and wetlands.  One study analysed water from 20 English rivers and found pesticide contamination of waterways via household drains, hypothesised to be from control of ectoparasites on pets. Seven out of 20 sites exceeded the level that would lead to chronic impact on ecosystems[21]

 

Herbicides

The use of glyphosate as herbicide varies within the environment and little is known about environmental concentrations in agricultural ponds and wetlands. The wide range of glyphosate compounds have different effects on amphibians; but the impact of direct over-spraying in terrestrial environments has been found to result in 79% mortality of Anurans[22]. Variation has been found from taxonomic group to taxonomic group with deformities appearing in larvae and one study reporting endocrine disruption, effects on development and inhibition of enzymes.  Other studies on the impact of ‘Roundup’ herbicide found that direct over-spray killed 96-100% of larval amphibians and 68-86% of post-metamorphic juvenile amphibians[23]. Additionally, herbicides have been found to indirectly increase amphibian susceptibility to trematode (parasite) infections by increasing the time spent in susceptible early developmental stages and suppressing tadpole immunity[24]. Research on common toad tadpoles found that younger tadpoles are more susceptible to glyphosate resulting in increased time to metamorphosis and reduced size of metamorphs[25]. Glyphosate has also been found to reduce tadpole survival and damage the cutaneous bacterial community[26]. Other studies have found that herbicide exposure can also disrupt amphibian immune responses1,[27], [28] and feminization of male gonadal development has been found following the use of the herbicide atrazine17.

 

Fertilisers

Changes in rainfall and runoff can lead to increased amounts of nitrogenous compounds and agricultural ponds and wetlands1. Ammonia and nitrite concentrations often increase after pollution incidents, but these break down to nitrates in waterbodies. A 2020 review of the impacts of nitrate on amphibians[29]  covered 46 studies and found nitrate led to a maximum decrease in activity of 79%, a decrease in growth of 29%, and can reduce survival by 62%. Nitrate exposure influenced all life stages except embryos, and nitrates in combination with other stressors can affect survival. Toxicology studies looking at ammonium nitrate[30]  and acidification of water bodies found that in the dwarf newt (a species in the same genus as the great crested newt) neither ammonium nitrate nor pH inhibited oviposition but the percentage of eggs that were ‘wrapped’ by females was lower at low pH, which could reduce survival rates.

 

Road de-icers

De-icing salts pose serious ecological risks to amphibians due to salinity increases and direct toxicity of chloride. Salts can have an extreme adverse effect on amphibians at all life stages[31], though embryonic and larval life stages are more sensitive than adults[32]. Road salt has been found to travel up to 172m from roads into wetlands and in the USA, egg mass densities of spotted salamanders and wood frogs were found to be reduced in roadside pools[33]. Road salts, as with many other pollutants, disrupt physiology and increase susceptibility to climate impact or disease. Populations exposed to salt runoff had slightly more frequent ranavirus related mass mortality events, more lethal infections, and greater pathogen environmental DNA[34]. Larvae raised in elevated salinity had 10 times more intense infections and shed five times as much virus as controls, suggesting increased transmission rates of viruses. In North America, wood frog populations living adjacent to roads had higher incidence and severity of oedema[35]  (bloating caused by subcutaneous fluid accumulation) during the breeding season, than frogs living away from the influence of roads. This was a direct impact of increased conductivity in breeding ponds, probably caused by road salt used for de-icing, and impacted movement, jumping ability and energy availability as well reducing muscle mass and causing skin damage.  Elevated salt concentrations have been found to increase both deformity frequency and severity32. Road salt (NaCl) in “low” concentrations has been shown to irreversibly disrupt the osmoregulatory ability of salamander eggs[36], [37] .

One study found that the concentration of total polycyclic aromatic hydrocarbons (PAH) from industrial waste, vehicle exhausts, oil leaks and sealants from pavement surfaces has increased while other persistent organic pollutants tended to remain stable or declined[38]. Urbanized watersheds experienced greater rises in PAH concentration compared to non-urban lakes with runoff from surfaces coated with coal–tar having higher concentrations of PAHs than runoff from asphalt-sealed or cement surfaces. Newts exposed to sediments containing coal–tar and asphalt had reduced ‘righting ability’ and diminished liver enzyme activities.

 

Metals

Some metals also have a sub-lethal effect on amphibians, raising their susceptibility to disease with an increase in evidence that lead accumulates in the tissues of fish and amphibians[39] and causes sub-lethal stress relating to haematology[40], oxidative stress, neurotoxicity and immune responses (in fish)[41] and affects metamorphosis and haematological[42],[43]and biochemical processes in amphibians[44].  The accumulation of lead in body tissues of these taxa will further impact predatory aquatic birds such as egrets, herons and kingfishers, feeding at the top of the food web. Heavy metal pollution effects movement and fitness of tadpoles[45] , with reduced swimming speed and disrupted swimming response suggesting a sub-lethal effect.  At a  UK Toad tunnel where amphibians cross below a busy road, seven out of eight tested metals including copper, zinc, lead and iron were in significantly greater concentrations in the tunnel than at a control site and were present in environmentally significant concentrations[46]  . Water samples also exhibited elevated concentrations of aluminium and chromium and occasionally an extreme alkaline pH, associated with leaching of portlandite in tunnel cements.

 

Microplastics

In 2019 the first record of microplastics from an amphibian stomach content (the stomach of the Italian crested newt) was recorded in a high-altitude environment in Italy[47]; an indication of the pervasive nature of microplastic pollution. Microplastics are transported by air or water and are small enough to be eaten by zooplankton and hence have a role within the food web in freshwater systems, where microplastic pollution maybe between 4 and 23 times higher than in the sea.[48].  In a 2022 review[49] investigating the impact of microplastics on amphibians found studies that indicated changes to body condition, increased susceptibility to infection or disease and where microplastics had an impact on survival and growth, bioaccumulation and toxic responses, malformation and swimming behaviour.

The extent of microplastic pollution is indicated by a 2020 study in Poland[50] which collected 201 tadpoles from 5 species at 8 different sites and found microplastics at all of the sites and in all five species; between them 26% of the tadpoles had ingested 71 different microplastic compounds. Links have been established between the number and variety of microplastics found within the larval stages of amphibians and the proximity of anthropogenic habitat to sites[51]. Where anthropogenic or ‘non-natural’ land cover was present the microplastic pollution in the larvae increased; roads have been found to be an important source of microplastic particles in the environment[52], through release of particles from tyres, road markings and polymer-modified bitumen, released through the wear and tear of tyres and the road surface.

 

Implications for amphibian metapopulations

In Scotland, a review of literature[53] found no Scottish study that has reported the effects of agrichemicals as demonstrated elsewhere, suggesting that much of the pollution is linked to agriculture that uses more agrochemicals. A 2021 study in Belgium[54]  sampled 26 amphibian breeding ponds and tested for 178 different compounds from 5 agrochemical groups to determine the frequencies of different compounds in wetlands.  Agrochemical compounds were present, with some occurring in only  1 of 26 ponds while others were in all 26. The 26 ponds were fed by runoff water and compounds found included anti-microbial drugs, anti-parasitic drugs, anthelmintics, mycotoxins, heavy metals and pesticides. 

In many amphibians, metapopulations consist of ‘source’ ponds and ‘sink’ ponds which vary year on year and in different environmental conditions.  Low annual survival of great crested newts (Triturus cristatus) is often related to mild winters and heavy rainfall, which can impact a metapopulation at the regional level[55]. A study in Kent found that some sub-populations suffered reproductive failure in most years, and recruitment to the metapopulation relied on the presence if a single ‘source’ pond. A population viability analysis at the Kent site confirmed that extinction risks increased when adult survival declined. In temperate ectotherms, mild temperatures during the winter can result in hibernating animals continuing to deplete energy reserves while being unable to feed, resulting in poor breeding condition the following spring. The ongoing sub-lethal effects of pollutants may act to exacerbate the impact of mild winters and increase the likelihood of an animals failing to breed or even failing to survive the winter.

Many of our amphibian species are in decline and based on the evidence presented above one could argue for a ‘rethink’ of our agricultural strategy to encourage the ‘re-construction’ of ponds for flood management and as reservoirs for biodiversity. The benefit of replacement of hedgerows and woodland to support pollinators and predators of insect pests will also benefit amphibians. Review of the use of pesticides and fertilisers and their impact on amphibians and on invertebrates, fish and animals further up the food chain will ensure there is minimal risk to humans and benefit to biodiversity.  This risk of acute or chronic toxicity to amphibian species from agrochemicals in our managed landscape is high and, along with:

  • loss of terrestrial habitat such as hedgerows and woodland,
  • loss of aquatic habitat with the ‘pondscape’ leading to reduction in pond density
  • road mortality during migration
  • invasive species and fish introductions
  • non-native diseases
  • the impact of climate change on overwintering
  • climate change leading to drought events

they cause sub-lethal effects that impact reproduction, metamorphosis and resistance to parasites and disease. Action is needed now to change the way agrochemicals are used, so we can avoid the loss of nocturnal amphibian choruses and a new ‘Silent Spring’.

 

References

[1] Mann, R.M., Hyne, R.V., Choung, C.B. and Wilson, S.P., 2009. Amphibians and agricultural chemicals: review of the risks in a complex environment. Environmental pollution, 157(11), pp.2903-2927.

[2] Egea‐Serrano, A., Relyea, R.A., Tejedo, M. and Torralva, M., 2012. Understanding of the impact of chemicals on amphibians: a meta‐analytic review. Ecology and evolution, 2(7), pp.1382-1397. 

[3] Köhler, H.R. and Triebskorn, R., 2013. Wildlife ecotoxicology of pesticides: can we track effects to the population level and beyond?. Science, 341(6147), pp.759-765.

[4]  Baker, N.J., Bancroft, B.A. and Garcia, T.S., 2013. A meta-analysis of the effects of pesticides and fertilizers on survival and growth of amphibians. Science of the total environment, 449, pp.150-156. 

[5] Smalling, K. L., Reeves, R., Muths, E., Vandever, M., Battaglin, W. A., Hladik, M. L., & Pierce, C. L. (2015). Pesticide concentrations in frog tissue and wetland habitats in a landscape dominated by agriculture. Science of the Total Environment, 502, 80-90.

[6]  de Oliveira, C.P.A., Carneiro, A.A., Ervilha, L.O.G., Machado-Neves, M., Souza, A.C.F. and Carvalho, R.P.R., Does environmental pollution affect male reproductive system in naturally exposed vertebrates? A systematic review. Theriogenology, 198, pp.305-316.

[7] Van Meter, R.J., Adelizzi, R., Glinski, D.A. and Henderson, W.M., 2019. Agrochemical mixtures and amphibians: the combined effects of pesticides and fertilizer on stress, acetylcholinesterase activity, and bioaccumulation in a terrestrial environment. Environmental toxicology and chemistry, 38(5), pp.1052-1061.

[8] Macdonald, K.J., Driscoll, D.A., Macdonald, K.J., Hradsky, B. and Doherty, T.S., 2023. Meta‐analysis reveals impacts of disturbance on reptile and amphibian body condition. Global Change Biology, 29(17), pp.4949-4965.

[9] Ji, C., Song, Q., Chen, Y., Zhou, Z., Wang, P., Liu, J., Sun, Z. and Zhao, M., 2020. The potential endocrine disruption of pesticide transformation products (TPs): The blind spot of pesticide risk assessment. Environment international, 137, p.105490.

[10] Denoël, M., Libon, S., Kestemont, P., Brasseur, C., Focant, J.F. and De Pauw, E., 2013. Effects of a sublethal pesticide exposure on locomotor behavior: a video-tracking analysis in larval amphibians. Chemosphere, 90(3), pp.945-951.

[11] Van Meter, R.J., Glinski, D.A., Purucker, S.T. and Henderson, W.M., 2018. Influence of exposure to pesticide mixtures on the metabolomic profile in post-metamorphic green frogs (Lithobates clamitans). Science of the Total environment, 624, pp.1348-1359.

[12] Agostini, M.G., Roesler, I., Bonetto, C., Ronco, A.E. and Bilenca, D., 2020. Pesticides in the real world: The consequences of GMO-based intensive agriculture on native amphibians. Biological Conservation, 241, p.108355.

[13] Woodley, S.K., Mattes, B.M., Yates, E.K. and Relyea, R.A., 2015. Exposure to sublethal concentrations of a pesticide or predator cues induces changes in brain architecture in larval amphibians. Oecologia, 179, pp.655-665.

[14] Capaldo, A., Gay, F., Valiante, S., De Falco, M., Sciarrillo, R., Maddaloni, M. and Laforgia, V., 2012. Endocrine-disrupting effects of nonylphenol in the newt, Triturus carnifex (Amphibia, Urodela). Comparative Biochemistry and Physiology Part C: Toxicology & Pharmacology, 155(2), pp.352-358.

[15] Bernabò, I., Biasone, P., Macirella, R., Tripepi, S. and Brunelli, E., 2014. Liver histology and ultrastructure of the Italian newt (Lissotriton italicus): Normal structure and modifications after acute exposure to nonylphenol ethoxylates. Experimental and Toxicologic Pathology, 66(9-10), pp.455-468.

[16] Nascimento, Í.F., Guimarães, A.T.B., Ribeiro, F., de Lima Rodrigues, A.S., Estrela, F.N., da Luz, T.M. and Malafaia, G., 2021. Polyethylene glycol acute and sub-lethal toxicity in neotropical Physalaemus cuvieri tadpoles (Anura, Leptodactylidae). Environmental Pollution, 283, p.117054

[17] Orton, F. and Tyler, C.R., 2015. Do hormone‐modulating chemicals impact on reproduction and development of wild amphibians?. Biological Reviews, 90(4), pp.1100-1117.

[18] Lürling, M. and Scheffer, M., 2007. Info-disruption: pollution and the transfer of chemical information between organisms. Trends in Ecology & Evolution, 22(7), pp.374-379.

[19] Brühl, C.A., Schmidt, T., Pieper, S. and Alscher, A., 2013. Terrestrial pesticide exposure of amphibians: an underestimated cause of global decline?. Scientific reports, 3(1), p.1135.

[20] Preston-Allen, R., Albini, D., Barron, L.., Collins, C.M., Dumbrell, A., Duncalf-Youngson, H., Jackson, M.., Johnson, A.., Prentis, A., Spurgeon, D. and Stasik, N., 2023. Are urban areas hotspots for pollution from pet parasiticides?.Imperial College London, Grantham institute Briefing Note No 15.

[21] Perkins, R., Whitehead, M., Civil, W. and Goulson, D., 2021. Potential role of veterinary flea products in widespread pesticide contamination of English rivers. Science of The Total Environment, 755, p.143560.

[22] Wagner, N., Reichenbecher, W., Teichmann, H., Tappeser, B. and Lötters, S., 2013. Questions concerning the potential impact of glyphosate‐based herbicides on amphibians. Environmental Toxicology and Chemistry, 32(8), pp.1688-1700.

[23] Relyea, R.A., 2005. The lethal impact of Roundup on aquatic and terrestrial amphibians. Ecological applications, 15(4), pp.1118-1124

[24] Rumschlag, S.L., Halstead, N.T., Hoverman, J.T., Raffel, T.R., Carrick, H.J., Hudson, P.J. and Rohr, J.R., 2019. Effects of pesticides on exposure and susceptibility to parasites can be generalised to pesticide class and type in aquatic communities. Ecology letters, 22(6), pp.962-972.

[25] Mikó, Z., Ujszegi, J. and Hettyey, A., 2017. Age-dependent changes in sensitivity to a pesticide in tadpoles of the common toad (Bufo bufo). Aquatic Toxicology, 187, pp.48-54.

[26] Krynak, K.L., Burke, D.J. and Benard, M.F., 2017. Rodeo™ herbicide negatively affects Blanchard’s cricket frogs (Acris blanchardi) survival and alters the skin-associated bacterial community. Journal of Herpetology, 51(3), pp.402-410.

[27] Rohr, J.R. and McCoy, K.A., 2010. A qualitative meta-analysis reveals consistent effects of atrazine on freshwater fish and amphibians. Environmental health perspectives, 118(1), pp.20-32.

[28] Knutie, S.A., Gabor, C.R., Kohl, K.D. and Rohr, J.R., 2018. Do host‐associated gut microbiota mediate the effect of an herbicide on disease risk in frogs?. journal of animal ecology, 87(2), pp.489-499.

[29] Isaza, D.F.G., Cramp, R.L. and Franklin, C.E., 2020. Living in polluted waters: A meta-analysis of the effects of nitrate and interactions with other environmental stressors on freshwater taxa. Environmental Pollution, 261, p.114091.

[30] Ortiz-Santaliestra, M.E., Marco, A., Fernández-Benéitez, M.J. and Lizana, M., 2007. Effects of ammonium nitrate exposure and water acidification on the dwarf newt: The protective effect of oviposition behaviour on embryonic survival. Aquatic toxicology, 85(4), pp.251-257.

[31] Hopkins, G.R., French, S.S. and Brodie Jr, E.D., 2013. Potential for local adaptation in response to an anthropogenic agent of selection: effects of road deicing salts on amphibian embryonic survival and development. Evolutionary Applications, 6(2), pp.384-392.

[32] Turtle, S.L., 2000. Embryonic survivorship of the spotted salamander (Ambystoma maculatum) in roadside and woodland vernal pools in southeastern New Hampshire. Journal of Herpetology, pp.60-67.

[33] Karraker, N.E., Gibbs, J.P. and Vonesh, J.R., 2008. Impacts of road deicing salt on the demography of vernal pool‐breeding amphibians. Ecological Applications, 18(3), pp.724-734.

[34] Hall, E.M., Brunner, J.L., Hutzenbiler, B. and Crespi, E.J., 2020. Salinity stress increases the severity of ranavirus epidemics in amphibian populations. Proceedings of the Royal Society B, 287(1926), p.20200062.

[35] Brady, S.P., Goedert, D., Frymus, L.E., Zamora‐Camacho, F.J., Smith, P.C., Zeiss, C.J., Comas, M., Abbott, T.A., Basu, S.P., DeAndressi, J.C. and Forgione, M.E., 2022. Salted roads lead to oedema and reduced locomotor function in amphibian populations. Freshwater Biology, 67(7), pp.1150-1161.

[36] Karraker, N.E. and Gibbs, J.P., 2011. Road deicing salt irreversibly disrupts osmoregulation of salamander egg clutches. Environmental Pollution, 159(3), pp.833-835.

[37] Karraker, N.E. and Gibbs, J.P., 2011. Contrasting road effect signals in reproduction of long-versus short-lived amphibians. Hydrobiologia, 664, pp.213-218.

[38] Bommarito, T., Sparling, D.W. and Halbrook, R.S., 2010. Toxicity of coal–tar and asphalt sealants to eastern newts, Notophthalmus viridescens. Chemosphere, 81(2), pp.187-193.

[39] Jofré, M.B., Antón, R.I. and Caviedes-Vidal, E., 2012. Lead and cadmium accumulation in anuran amphibians of a permanent water body in arid Midwestern Argentina. Environmental Science and Pollution Research, 19, pp.2889-2897.

[40] Tewari, H., Gill, T.S. and Pant, J., 1987. Impact of chronic lead poisoning on the hematological and biochemical profiles of a fish, Barbus conchonius(Ham). Bulletin of environmental contamination and toxicology, 38(5), pp.748-752.

[41] Lee, J.W., Choi, H., Hwang, U.K., Kang, J.C., Kang, Y.J., Kim, K.I. and Kim, J.H., 2019. Toxic effects of lead exposure on bioaccumulation, oxidative stress, neurotoxicity, and immune responses in fish: A review. Environmental toxicology and pharmacology, 68, pp.101-108.

[42] Fink, N. and Salibián, A., 2005. Toxicological studies in adult amphibians: effects of lead. Applied Herpetology, 2(3), pp.311-333.

[43] Rosenberg, C.E., Peri, S.I., Arrieta, M.A., Fink, N.E. and Salibián, A., 1998. Red blood cell osmotic fragility in Bufo arenarum exposed to lead. Archives of physiology and biochemistry, 106(1), pp.19-24.

[44] Arrieta, M.A., Bruzzone, L., Apartín, C., Rosenberg, C.E., Fink, N.E. and Salibián, A., 2004. Biosensors of inorganic lead exposure and effect in an adult amphibian. Archives of environmental contamination and toxicology, 46, pp.224-230

[45] Zhang, W., Zhi, H. and Sun, H., 2020. Effects of heavy metal pollution on fitness and swimming performance of Bufo raddei tadpole. Bulletin of Environmental Contamination and Toxicology, 105, pp.387-392.

[46] White, K.J., Mayes, W.M. and Petrovan, S.O., 2017. Identifying pathways of exposure to highway pollutants in great crested newt (Triturus cristatus) road mitigation tunnels. Water and Environment Journal, 31(3), pp.310-316.

[47] Lannella, M., Console, G., D’Alessandro, P., Cerasoli, F., Mantoni, C., Ruggieri, F., Di Donato, F. and Biondi, M., 2019. Preliminary analysis of the diet of Triturus carnifex and pollution in mountain Karst ponds in central Apennines. Water, 12(1), p.44.

[48] Horton, A.A., Walton, A., Spurgeon, D.J., Lahive, E. and Svendsen, C., 2017. Microplastics in freshwater and terrestrial environments: Evaluating the current understanding to identify the knowledge gaps and future research priorities. Science of the total environment, 586, pp.127-141.

[49] Burgos-Aceves, M.A., Faggio, C., Betancourt-Lozano, M., González-Mille, D.J. and Ilizaliturri-Hernández, C.A., 2022. Ecotoxicological perspectives of microplastic pollution in amphibians. Journal of Toxicology and Environmental Health, Part B, 25(8), pp.405-421.

[50] Kolenda, K., Kuśmierek, N. and Pstrowska, K., 2020. Microplastic ingestion by tadpoles of pond-breeding amphibians—first results from Central Europe (SW Poland). Environmental Science and Pollution Research, 27(26), pp.33380-33384.

[51] Szkudlarek, M., Najbar, B. and Jankowiak, Ł., 2023. Microplastics pollution in larvae of toads, frogs and newts in anthropopressure gradient. Ecological Indicators, 155, p.110971.

[52] Rødland, E.S., Okoffo, E.D., Rauert, C., Heier, L.S., Lind, O.C., Reid, M., Thomas, K.V. and Meland, S., 2020. Road de-icing salt: assessment of a potential new source and pathway of microplastics particles from roads. Science of the Total Environment, 738, p.139352.

[53] Downie, J.R., Larcombe, V. and Stead, J., 2019. Amphibian conservation in Scotland: A review of threats and opportunities. Aquatic Conservation: Marine and Freshwater Ecosystems, 29(4), pp.647-654.

[54] Goessens, T., De Baere, S., Deknock, A., De Troyer, N., Van Leeuwenberg, R., Martel, A., Pasmans, F., Goethals, P., Lens, L., Spanoghe, P. and Vanhaecke, L., 2022. Agricultural contaminants in amphibian breeding ponds: Occurrence, risk and correlation with agricultural land use. Science of the Total Environment, 806, p.150661.

[55] Griffiths, R.A., Sewell, D. and McCrea, R.S., 2010. Dynamics of a declining amphibian metapopulation: survival, dispersal and the impact of climate. Biological Conservation, 143(2), pp.485-491.

 

 

Croaking Science- Another amphibian reproductive mode: newly-hatched caecilians feed on mother’s milk.

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Written by Roger Downie, Froglife and University of Glasgow

Although we mostly think of amphibians mating in water, depositing their eggs there, and leaving them to develop unaided, the group actually includes a vast diversity of reproductive modes, 74 at the last comprehensive count (Nunes-de-Almeida et al., 2021), many of them including some kind of parental care. Now comes something new: Mailho-Fontana et al.(2024) report that in Siphonops annulatus, a widely-distributed South American caecilian, mothers produce and release a milk to feed their newly-hatched babies.

Caecilian babies around the mother’s vent. Credit: Carlos Jared

The 220+ species in the order Gymnophiona, commonly called caecilians, are acknowledged as the most poorly known amphibian group, a situation related to their tropical distribution and secretive burrowing habits. One feature that is known about the group is that a relatively high proportion of species (about 20%) is viviparous (giving birth to live young), compared to the tiny proportion of anurans and urodeles that reproduce in that way. In viviparous caecilians, the eggs are retained in the oviduct and hatch there, with the young nourished by oviductal secretions, growing substantially during a gestation period that can be as long as 11 months.

Egg-laying caecilians also show parental care, with mothers often looking after the eggs, and at least in some cases, feeding the hatchlings on modified, lipid-rich skin which the young browse on using specialised teeth. This behaviour has previously been reported in S. annulatus by Wilkinson et al. (2008).

Nunes-de-Almeida and colleagues were able to capture and observe for an extended period in captivity a sample of 16 adult female S. annulatus. After hatching, the young (4-13 per clutch) clustered around the coiled-up mother, keeping close to her vent. Periodically, the young touched the vent and emitted high-pitched sounds. These behaviours seemed to elicit the release of a transparent, viscous milk, which the young fed on. After feeding, the young became relatively inactive for a time. The young may compete for access to the milk, since data showed considerable variation in growth rate increase among a batch of young; on average, body mass increased by 130% during the week after hatching. Milk release was at least daily, and up to six times a day. As in Wilkinson et al.’s study, skin feeding also occurred, but less frequently, about once a week. Histological analysis of the female reproductive tract showed that milk is produced by oviductal glands which develop during the reproductive period. These are very similar to the glands which appear during gestation in viviparous species, but have not previously been reported in egg-layers. The lipid content of the milk is mainly composed of long-chain fatty acids, palmitic and stearic; the milk is also rich in proteins and carbohydrates. The period of maternal care, during which they do not feed,  lasts about two months, during which maternal body mass declines by 30%, so this behaviour is very costly to the mothers.

Milk production is a diagnostic feature of mammalian reproduction, but it does occur occasionally elsewhere in the animal kingdom: in pigeons and flamingoes among the birds, for example, where the crop produces a nutritive secretion. It will be fascinating to discover if S. annulatus is an isolated case among the caecilians. Frost (2024) lists four species in the genus Siphonops, with five genera and 28 species in the family Siphonopidae, at least some of which are known to show skin-feeding. In my view, it would be surprising if this combination of hatchling behaviour and maternal physiology  is restricted to a single species.

 

Acknowledgement I thank Malcolm Kennedy for drawing my attention to this fascinating discovery.

 

References

Frost (2024). Amphibian Species of the World. Accessed 15/3/24.

Nunes-de-Almeida et al. (2024). Milk provisioning in oviparous caecilian amphibians. Science 383, 1092-1095.

Wilkinson et al. (2008). One hundred million years of skin feeding? Extended parental care in a neotropical caecilian. Biology Letters 4, 358-361.

Wilkinson et al. (2013). A new species of skin-feeding caecilian, and the first report of the reproductive mode in Microcaecilia. PLOS ONE 8, e57756.

 

Croaking Science: are our pet’s flea treatments harming pondlife?

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Written by Elsbeth Leighton, Coalface to Wildspace Project Assistant

As a nation, the UK has a lot of pets, so much so that we are often labelled as a “nation of animal lovers”. Over half of us have pets, which includes 12 million dogs, 11 million cats and another few millions small animals such as rabbits and hamsters (Kantar/Soulor Consulting, 2023). Being responsible and caring pet owners, we want to do what’s best for our animals and follow vet advice about how to keep them healthy and happy. For many of us, that now includes a regular flea and worm treatment for our cats and dogs to prevent any nasty infestations.

This treating of healthy pets with regular doses of parasiticides, i.e. medicines to treat fleas, worms and other parasite infections, as a way to prevent rather than cure infestations is called prophylactic treatment as opposed to reactive treatment. This approach has the benefit that, since it can prevent infestations before they start, it also has a good chance of preventing the unpleasant complications that can sometimes arise from pet parasite infections, such as such as flea allergy dermatitis or disease spread (Murphy and Wright, 2020). This seems quite sensible, but could this approach be harming the environment and our wildlife?

It’s well known by now that pesticides used on crop fields have negative effects on wildlife and the environment (e.g. Sánchez-Bayo, 2021). Neonicotinoids, in particular, are now notorious for their deadly impact on insects and bees in particular, which earned them a ban from use in agriculture by the European Union in 2018 (European Commission, 2023). However, what many of us may not know is that many of those same pesticides that can no longer be used in agriculture are still allowed to be used in other settings, including in pet parasiticides. This decision has been justified by the reasoning that so little of these pesticides is used per pet and even less will reach the outside environment, so the threat to the environment must be very small and not worth worrying about.

However, a recent review of the research around pet parasiticides by Preston-Allen and their team from the Grantham Institute, shows that this assumption is incorrect and our fleaing and worming treatments present a real danger to wildlife, and freshwater habitats in particular. They point out that imidacloprid, one of the most popular pet parasiticides as well as a neonicotinoid banned for agricultural use, was found at toxic levels in two thirds UK rivers surveyed in a recent study (Perkins et al, 2021). Given that these chemicals are no longer used in agriculture, the most likely source is from our pets. Research looking into how pet parasiticides might get into our freshwater habitats indicated that wastewater, i.e from our drains and our washing machines, is the main route, but that shedding of pet hair, pet urine and outdoor swimming are also causes (Preston-Allen et al, 2023, Diepens et al, 2023).

Once they have made it into the environment and into our ponds, lakes and rivers, these chemicals can have devastating impacts on wildlife. These parasiticides are designed to kill invertebrates and this includes our freshwater insects. The research shows that they are deadly to many species of freshwater invertebrates, and they can also have serious negative effects on other animals, including fish, birds and mammals (Preston-Allen et al, 2023). Not all of these animals die from the parasiticides, but they may be impacted in other ways. For example, one study showed that fish with pet parasiticides in their water swim slower and lower in the water, which makes it harder for them to forage for food and easier for predators to catch them (Domingues et al, 2016). Effects like these can impact on the relationships between species and damage the overall health of our ecosystems.

Not only does the current prophylactic approach to pet parasiticides pose a threat to the environment and our wildlife, but there are also concerns it will lead to parasites developing resistance to these drugs (Nind and Mosedale, 2022). This is analogous to the issue of antibiotic resistance, where overuse of some drugs leads to the diseases that we want to target becoming resistant to antibiotics due to overexposure, making it harder to treat these infections (e.g. GLASS Report, 2022). If we keep overusing these pet parasiticides, it is highly likely that these parasites we are aiming to prevent will become resistant and treating infestations in the future will be much more challenging.  

So, what can we as pet owners do?

The key recommendation from the researchers is to start treating these issues reactively instead of prophylactically if you can. This means checking our pets regularly for any signs they may have fleas or worms and seeking vet treatment when it arises, instead of regularly dosing healthy pets with parasiticides. This might not be the ideal solution for everyone as the risks from infestations are higher for some people, such as those with pets that are more vulnerable to complications from fleas or owners who are immunocompromised. So, we should all individually evaluate the level of risk we and our pets face to determine the best approach for us that balances our needs against the environmental impact of these parasiticides.   

As well as aiming to limit the amount of parasiticide treatments we use through a return to reactive treatment, we can aim to reduce the amount of these chemicals that leak into the environment when we do use them. This might mean keeping your cat inside for a few days after applying a spot-on treatment, or not washing your pets bedding as frequently when your pets have been treated, as these are both ways parasiticides can make their way into the environment (Preston-Allen et al, 2023). Another key step we should all try to take is being careful to keep our dogs out of streams and other water bodies when they’ve been treated for parasites recently. It is best to try to avoid letting dogs in wild water bodies in general, as they can stir up and disturb the organisms and plants living there, but especially when they have just been treated for parasiticides as they can wash off into the water (froglife, 2022).

 

References

Diepens, N.J., Belgers, D., Buijse, L., Roessink, I., (2023) Pet dogs transfer veterinary medicines to the environment. Science of The Total Environment. https://doi.org/10.1016/j.scitotenv.2022.159550.

Domingues, I., Oliveira, R., Soares, Amorim, M.V.M., et al. (2016). Effects of ivermectin on Danio rerio: a multiple endpoint approach: behaviour, weight and subcellular markers. Ecotoxicology. https://doi.org/10.1007/s10646-015-1607-5.

European Commission, (2023). Neonicitinoids, https://food.ec.europa.eu/plants/pesticides/approval-active-substances/renewal-approval/neonicotinoids_en,  accessed 15/12/2023

Froglife, (2022). #pawsagainstponds Campaign, https://www.froglife.org/2022/03/22/pawsagainstponds-campaign/,  accessed 15/12/2023

Global antimicrobial resistance and use surveillance system (GLASS) report, (2022). Geneva: World Health Organization. Licence: CC BY-NC-SA 3.0 IGO.

Kantar/Soulor Consulting, (2023). online survey with 8,901 respondents, https://www.ukpetfood.org/information-centre/statistics/uk-pet-population.html

Murphy, D, Wright, I., (2020). Value of using preventive pet parasiticides. The Veterinary Record. Feb 29;186(8):252. doi: 10.1136/vr.m727.

Nind, F., Mosedale, P., (2022). Antiparasitic Resistance. BSAVA Guide To The Use Of Veterinary Medicines. https://www.bsavalibrary.com/content/chapter/10.22233/9781905319862.chap20?crawler=true&mimetype=application/pdf

Perkins, R., Whitehead, M., Civil, W., Goulson, D., (2021). Potential role of veterinary flea products in widespread pesticide contamination of English rivers. Science of The Total Environment. https://doi.org/10.1016/j.scitotenv.2020.143560.

Preston-Allen, R.G.G., Albini, D., Barron, L., Collins, T., Dumbrell, A., Duncalf-Youngson, H., Jackson, M., Johnson, A., Perkins, R., Prentis, A., Spurgeon, D., Stasik, N., Wells, C. and Woodward, G., (2023). Are urban areas hotspots for pollution from pet parasiticides? Grantham Institute Briefing. note #15. Doi: https://doi.org/10.25561/102699

Sánchez-Bayo F., (2021). Indirect Effect of Pesticides on Insects and Other Arthropods. Toxics. Jul 30;9(8):177. doi: 10.3390/toxics9080177.

Croaking Science: Portuguese man-o-war and leatherback turtle wash up in Cornwall

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As things calm down after the recent storms, Cornwall has had a number of interesting strandings: in mid-October there were reports of Portuguese Man-O-war jellyfish stranded on beaches near Porthleven. 

Froglife Conservation & Science Manager, Andrew Smart, came across them on nearby Loe Bar.  “I think we probably saw about 10 gas floats along about 500m of beach; they were still really pink so they’d not been on the shore long; you could see the blue base where the trailing tentacles would have attached.” 

The Portuguese man-o-war isn’t actually a jellyfish, but a colonial organism made up of individual ‘zooids’ that have different functions in the colony, such as digestion and stinging tentacles. Recently there have been more regular strandings along the coasts of Cornwall. The gas filled floats are caught by the wind and the man-o-war floats across the ocean following winds and currents, trailing its stinging tentacles behind it.

The oceanic predators of Portuguese man-o-war include the sunfish, which also occurs off Cornwall, and sea turtles, including the UK’s largest reptile, the leatherback turtle. The leatherback is a member of the British Fauna and a UK protected species, with animals in UK waters migrating between the UK and the Caribbean Sea where they nest (see our Croaking Science article here for more information)

Last week, a dead leatherback turtle washed up on Perranporth beach in North Cornwall It was quite badly decomposed and was collected by the Marine Strandings Network and taken away for further analysis. Often leatherbacks die because they swallow floating plastic bags they mistake as jellyfish and investigation of the dead animal will help us understand the extent of this problem. 

This animal measured 1.66m, which sounds large, but the largest recorded, washed up in Wales in the 1908s, was measured at 2.9m (9½ ft) long!  Its quite late in the year for a leatherback stranding but the sea temperature this year off Cornwall is around 13-15  ͦC, higher than the average for November (11- 12  ͦC) and warmer temperatures may have resulted in more jellyfish and salps for leatherbacks to feed on.

At the start of 2023 there were a high number of strandings of other species, small loggerheads and Kemps ridley’s. Sadly, many of these small turtles die from cold shock because they are not adapted to our colder waters like the leatherback, but if they are found quickly enough they can be nursed back to health and released.

Leatherback turtles are stranded and sighted off our coasts every year:

Year

Strandings & sightings

2018

17

2019

15

2020

9

2021

17

2022

9

Data from Marine Environmental Monitoring Turtle Strandings & Sightings Annual Reports 2018-2022

If you’re out walking our western beaches keep an eye out for stranded turtles. They can be stranded anywhere although mainly on the west and south coasts, as can be seen from this diagram of strandings and sightings over the last 10 years, recorded by R.S. Penrose and M.J.B. Westfield in the May 2023 Marine Environmental Monitoring British & Irish Marine Turtle Strandings & Sightings Annual Report 2022

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