U.S. Geological Survey accepted manuscript accepted manuscript AM U.S. Department of the Interior U.S. Geological Survey 70274234 70274234Chronic Exposure to Waterborne Nickel Significantly Reduced Growth of Juvenile Crayfish (Faxonius virilis Adrian P. Moore1, Mark L. Wildhaber1, Zachary D. Beaman1, Kendell R. Bennett2, Karlie K. Ditter1, Danielle Cleveland1, Jenna A. Blanton1, Tyler J. Grant1

Corresponding Author: Adrian P. Moore; U.S. Geological Survey, Columbia Environmental Research Center. 4200 New Haven Road, Columbia, MO 65201, USA. Email: amoore@usgs.gov

U.S. Geological Survey (USGS), Columbia Environmental Research Center, 4200 New Haven Road, Columbia, MO 65201, USA

U.S. Fish and Wildlife Service (USFWS), Southwestern Native Aquatic Resources and Recovery Center, 7116 Hatchery Road, Dexter, NM 88230, USA

 2026 U.S. Geological Survey Reston, Virginia Abstract

Crayfish are critical functional components of aquatic ecosystems. Previous research has documented adverse effects of mineral extraction on crayfish. Here, we characterize potential risks of mining-derived waterborne nickel (Ni) to crayfish by documenting the effects of dissolved Ni on growth and food consumption of juvenile virile crayfish (Faxonius virilis) in a 28-day chronic laboratory exposure. Nominal Ni concentrations ranged from 31.25 to 500 micrograms per liter (µg/L; pH = 7.96 ± 0.20, hardness = 150 ± 1 milligrams per liter as calcium carbonate). Crayfish survival, carapace length, and wet weight were measured. After 28 days of exposure, a 24-h feeding trial was performed to determine differences in food consumption. During the growth trial, 99% of crayfish survived. Change in wet weight and final wet weight were the most sensitive endpoints, with 20% effect concentrations of 24.8 and 22.6 µg/L Ni, respectively. Crayfish exposed to an average of 438 µg/L Ni consumed 41% less, and weighed 65.1% less, than control crayfish. These results suggest chronic, sublethal exposure to waterborne Ni may have negative effects on crayfish growth. Reduced growth and consumption rates in crayfish could have wide-ranging consequences throughout aquatic ecosystems since crayfish are consumers, prey, keystone trophic regulators, and ecosystem engineers. Finally, these results could inform bioenergetics and may be coupled with population models to predict potential changes in population sizes of native and invasive crayfishes.

Keywords

Crayfish, Metal toxicity, Growth rates, Sub-lethal exposure, Critical minerals

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 Introduction

Metal pollution is a global issue posing risks to human and wildlife health (Su et al. 2014). Anthropogenic activities have increased concentrations of metals released into the environment worldwide, leading to the accumulation of metals in both terrestrial and aquatic systems (Järup 2003; Zhou et al. 2020). Metal pollutants can alter ecosystem function and may enhance invasion potential by nonnative species via reduction of native species biodiversity; pollution may also create favorable conditions for establishment of invasive species (Capinha et al. 2013; Sun et al. 2023). For example, copper (Cu) was found to reduce species richness of native marine invertebrates on fouling panels in an estuarine environment, while richness of exotic species did not differ significantly between control and Cu treatments (Crooks et al. 2011). One of the primary sources of metal pollution in aquatic systems worldwide is the mining lifecycle, which includes historical waste piles, active extraction, and mineral processing activities. One such metal pollutant is nickel (Ni), which is an essential element for aquatic plants and cyanobacteria in trace amounts (at ≤0.005 µM (micromolar) for many aquatic plants; Muyssen et al. 2004) but becomes toxic to plants and animals at greater concentrations (Binet et al. 2017). Nickel occurs naturally in the environment and is released by weathering of exposed rock or by volcanic eruptions. It is a critical mineral under U.S. 87 Federal Register 10381 (U.S. Geological Survey 2025) and is widely used in industrial materials and processes (Shukla et al. 2001; Ni Institute 2021). Nickel can be released into the atmosphere through coal and oil combustion and into soil and water through mining, smelting, and other processes (Genchi et al. 2020). The U.S. Environmental Protection Agency (EPA) freshwater Criterion Continuous Concentration for Ni is 52 µg/L (at a hardness of 100 mg/L as calcium carbonate [CaCO3]) for protection of aquatic life (USEPA 2025). Once released into the environment, wind, rain, and fire events can move contaminated wastes across the landscape (Lawlor and Tipping 2003; Doig and Liber 2006). With applications of Ni increasing, there is concern that significant amounts will be released into the environment (El-Naggar et al. 2021). The European Union and Canada consider Ni a priority substance—a substance that poses a threat to human health or the environment and is targeted for reduction and/or risk management (European Parliament and Council of the European Union, 2013; Government of Canada, 1999). For these reasons, it is important to understand the ecological consequences of environmental Ni.

Crayfish are integral components of aquatic ecosystems throughout the world, acting as consumers, prey, keystone trophic regulators, and ecosystem engineers (Reynolds et al. 2013). As keystone consumers and opportunistic omnivores, crayfish engage with the food web at multiple levels (Taylor and Soucek 2010; Gherardi et al. 2011); moreover, they are often the largest and longest-lived macroinvertebrate in aquatic systems (Gherardi et al. 2011; Allert et al. 2012). Crayfish are generally exposed to and uptake metals from surface and pore waters, sediments, and their food due to their benthic lifestyle and interaction with sediment, sediment-contaminated detritus, and other habitat-derived items (Velez-Gavilan 2016; Allert et al. 2024). Sediments often serve as a sink for metals, and concentrations temporally flux at the water-sediment interface in pore water and labile sediments where crayfish and other benthic invertebrates dwell (Amato et al. 2015). In this way, crayfish are vulnerable to the toxic effects of metals pollution and have been previously studied for use as bioindicators of degraded water quality or habitat (Allert et al. 2012; Reynolds et al. 2013; Allert et al. 2024). While metals contamination may impair native crayfish populations, it may also promote the invasion of nonnative species. Invasion by nonnative crayfish and the subsequent effects on native species populations are of growing concern (Gherardi et al. 2011), so it is important to understand how metals contamination may exacerbate these concerns in sensitive areas (Svobodová et al. 2012).

The few studies that have documented the effects of aquatic metal contamination on crayfish have shown negative physiological effects. Metals such as Cu, cadmium (Cd), lead (Pb), and zinc (Zn) have been shown to be toxic to juvenile calico crayfish (Faxonius immunis) via inhibition of oxygen consumption (Khan et al. 2006). Investigation of a mass mortality event of noble crayfish (Astacus astacus) and stone crayfish (Austropotamobius torrentium) in the Czech Republic indicated that the crayfish had experienced chronic exposure to aluminum (Al), iron (Fe), arsenic (As), Cd, Pb, Cu, Zn, and mercury (Hg; Svobodová et al. 2017). In Danube crayfish (Pontastacus leptodactylus), sublethal exposure to 17.3 milligrams per liter (mg/L) Ni (from the salt nickel chloride hexahydrate) led to the accumulation of Ni in germ and somatic cells, resulting in physiological alterations in the reproductive system (Zarnescu et al. 2017). Moreover, metals in the water column are known to accumulate in the tissues and bind directly to the exoskeleton of crayfish (Devi et al. 1996; Kouba 2009; Simon and Stewart 2014).

In Missouri, the Ozark region is of special ecological concern due to the historical mining operations that took place in the Tri-State Mining District (TSMD; in Missouri, Kansas, and Oklahoma) and the Southeast Missouri Lead Mining District (SEMO-LMD), combined with the presence of numerous species of concern, including crayfish. The TSMD and SEMO-LMD regions were heavily mined for Pb and Zn from the early 1800s to the 1960s, and elevated metals concentrations are still present in the riparian habitats (Allert et al. 2012) due to continued releases from historical waste piles. The Ozark region is also home to the federally listed, endangered Benton County cave crayfish (Cambarus aculabrum), the Big Creek crayfish (Faxonius peruncus) and the threatened St. Francis River crayfish (Faxonius quadruncus; U.S. Fish and Wildlife Service 2025). Since crayfish are the most abundant macroinvertebrate in many streams (Rabeni 1985), they play an important role in the stream communities of this area. Understanding the effects of differing concentrations of metal contaminants on the survival and life history of crayfish is critical to stream conservation efforts. Given the history of mining pollution in the Ozark region, coupled with crayfish being an integral part of that ecosystem, further research is needed to understand potential long-term effects of metals pollution on crayfish populations.

Metal pollution has been shown to adversely affect populations of crayfish in Missouri. Mining operations in the SEMO-LMD have led to the contamination of streams by Pb, Zn, cobalt (Co), Ni, and other metals, which co-occur in the target ores; concentrations of these metals exceed toxicity thresholds for aquatic invertebrates (Besser et al. 2009). Previous studies have found that concentrations of Pb, Cd, Zn, and other metals are negatively correlated with crayfish density and that crayfish survival is lower downstream of former mining sites (Allert et al. 2008; Allert et al. 2012; Allert et al. 2024). In a study measuring in situ survival of caged woodland crayfish (Faxonius hylas), crayfish survival and biomass were significantly lower at mining sites compared to reference sites (Allert et al. 2009). The St. Francis River crayfish was found to have reduced densities at sites downstream of mining-related metal releases from the Madison County Mines National Priority List Superfund site (MCM; within SEMO-LMD) compared to upstream sites that were not affected by mining wastes (Allert et al. 2024). Water, sediments, detritus, plant materials, and crayfish tissues at the downstream sites were found to be contaminated with Ni, Cd, Cu, Co, Pb, and Zn. Additionally, juvenile St. Francis River crayfish had decreased survival and growth in in situ cage and laboratory studies where crayfish were exposed to metal mixtures in MCM waters (Allert et al. 2024). The Ni concentrations to which crayfish were exposed varied by compartment (sediment, water, food). For example, the median concentration of Ni in surface water was 1.08 µg/L at the control site (upstream of known mining releases) while the sites affected by mining had medians ranging from 4 to 37 µg/L Ni (Allert et al. 2024). Median concentrations of Ni in pore water were 2.54 µg/L at the control site and 4–40 µg/L at mining-affected sites (Allert et al. 2024). In detritus, median concentrations of Ni were 23.1 µg/g dry weight at the control site and up to 329 µg/g dry weight at mining-affected sites. The maximum value recorded of Ni in detritus was 469 µg/g dry weight (Allert et al. 2024). The MCM site was recently re-permitted for extraction of Co and for processing ores for critical minerals, including Ni (Missouri Department of Natural Resources, 2024); in this way, the MCM remains an area of ecological concern in Missouri and the Ozarks region.

We chose the virile crayfish (Faxonius virilis) as our study species due to its potential to be exposed to metals contamination and its status as both an important native species and an invasive species, depending upon the location. In their native habitats, the virile crayfish is an important component of aquatic food webs. Virile crayfish consume aquatic macrophytes, macroinvertebrates, fish eggs, small amphibians, carrion, and detritus (Momot 1967). In turn, they are consumed by fish, otters, turtles, birds, snakes, and humans (NEMESIS 2023), making them vital for energy transfer among trophic levels. Virile crayfish have significant potential to be exposed to sediment-borne metals. They are not primary burrowers, but they will construct burrows when there is a lack of shelter, a high density of individuals, or in winter to avoid freezing temperatures (Caldwell and Richard, 1969; Peeters et al. 2024). Additionally, although the virile crayfish is native to much of the central United States, it has also established itself as an invasive species in parts of Europe and other regions of the United States (Durland Donahou 2025). While native to much of Missouri, including some parts of the Ozark Mountains, the virile crayfish is considered invasive in other parts of the Missouri Ozark Mountains (Durland Donahou 2025).

The overall goal of this study was to determine whether Ni contamination adversely affects crayfish growth and consumption in a laboratory setting, with the ultimate intent to potentially extrapolate to population level effects. We examined the effects of varying waterborne Ni concentrations on the growth (body weight) and consumption rates of virile crayfish in a laboratory setting. The laboratory setting was selected to allow for control of environmental factors-particularly to limit exposure to other metals and co-contaminants. Nickel concentrations selected in this study were determined using concentrations measured in a previous Missouri field study in MCM (Allert et al. 2024). We also aimed to collect data that could be used to develop quantitative relationships among Ni exposure, consumption, and growth; such datasets could be used in future studies incorporating metals effects in crayfish bioenergetics and population models. Thus, a better understanding of the effects of Ni pollution on virile crayfish could provide insight into potential effects on aquatic communities, food webs, and native and invasive species populations.

 Materials and Methods Study animals

The test organisms were juvenile virile crayfish (hereafter “crayfish” in the Materials and Methods and Results) of age 0 to 1 years. They were cultured in an outdoor culture pond filled with well water (hardness approximately 300 mg/L as CaCO3) at the U.S. Geological Survey’s (USGS) Columbia Environmental Research Center (CERC, Columbia, Missouri, USA). Juvenile crayfish were selected for this study because the juvenile life stage is often more sensitive than the adult life stage (Hutchinson et al. 1998; Wiggington and Birge 2007). Additionally, we sought to minimize biological variability by using similarly aged, pond-reared, young-of-year crayfish that resulted from a synchronous breeding event. Crayfish were retrieved from the culture pond and placed into aerated coolers filled with pond water, then acclimated to laboratory temperature over the course of 2–3 hours (h) by slowly mixing fresh well water with the culture pond water in the coolers. This first acclimation was intended to reduce any shock induced by adding crayfish to colder laboratory water (22 °C) from the warmer (23 °C) pond water. Once the organisms were in laboratory-temperature water, they were transferred to and held in a 726.8 liter (L) recirculating system with well water at a temperature of 22.2 ± 1.0 degrees Celsius (°C) for 3 weeks; the water turnover rate was 1.3 liters per minute (L/min). Approximately 250 crayfish were kept in the recirculating tank without sediment; adequate numbers of PVC shelters were provided to help minimize cannibalism. A cover was placed over the tank to keep crayfish from escaping. All crayfish were exposed to a light:dark cycle of 16:8 h. Four days prior to the start of the chronic exposures, the crayfish were transitioned to “base water” (well water diluted with deionized water to a hardness of 150 mg/L as CaCO3; alkalinity, 126 mg/L as CaCO3; pH, 7.96) by slowly exchanging well water for base water via the recirculating system (see Exposure system below). The base water hardness was based on hardness levels in SEMO-LMD streams (110–300 mg/L as CaCO3; Allert et al. 2024). Hardness and pH levels can affect toxicity of Ni (Pyle et al. 2002), so it was monitored and kept consistent and comparable to Missouri streams throughout the study. Before the experiment began, crayfish were fed a mixture of three sinking pellet feed: Otohime C2 Marine Larval and Grow-Out Feed (840–1,410 micrometers [µm]), Otohime S2 Marine Larval and Weaning Feed (1,400 µm), and Otohime EP1 (1.7 millimeters [mm]) Marine Larval and Grow-Out Feed (Reed Mariculture, Campbell, CA). They were acclimated to this food for a period of three weeks, then were fed exclusively EP1 pellets three days before the start of the study and throughout the study.

 Exposure system

Exposures occurred in two intermittent proportional flow-through diluter systems, based on Mount and Brungs (1967) and more fully described by Brunson et al. (1998; Fig. 1). Each diluter system included a temperature-controlled water bath (mean ± 1 SD = 21.1 ± 0.2 °C). Twelve 75.7-L aquaria were connected to the diluter system within each water bath (Fig. 1a). There was a total of 24 aquaria across the two diluters, with one aquarium representing an experimental unit. There were four replicate aquaria for each treatment (two per diluter), each containing four exposure chambers per aquarium that were duplicates, and one crayfish per exposure chamber (i.e., N = 96 crayfish total). In this way, there were n = 16 crayfish per Ni concentration. Each aquarium was equipped with four nearly identical “exposure chambers,” with one exposure chamber representing a subsample, to separate individual crayfish from each other during a trial (Fig. 1a); this separation prevented antagonistic behaviors among individuals. Dividers made of opaque white polypropylene sheeting were placed between the four exposure chambers to provide a visual barrier between exposure chambers. Each aquarium served as a water bath for its four exposure chambers and was equipped with a standpipe to maintain a constant water level. Exposure chambers were made from 1-L glass beakers that had been modified by removing rectangular section of glass (2.5 centimeters [cm] by 5.7-cm) from the top rim of the beaker. This removed section was replaced with mesh stainless-steel screen to allow exposure water replacement. The diluter systems delivered approximately 1 L of additions of fresh base control water or base water containing Ni (collectively “treatment solutions”) once every hour. Fresh treatment solution was divided evenly among the four exposure chambers. At each cycle of water addition, excess water in the exposure chambers flowed out through the screen, with no water exchange among exposure chambers (Fig. 1a). Between hourly additions of fresh treatment solutions, the water level in each aquarium was maintained below the mesh screen using the aquarium’s standpipe. Each exposure chamber held approximately 950 milliliters (mL) of treatment solution. Each exposure chamber was equipped with an air line to provide gentle aeration. Nylon mesh screens attached to plastic grates were placed over the tops of each set of exposure chambers to prevent escape of crayfish without impeding flow of incoming treatment solution.

During experimental trials, a treatment solution was automatically delivered to the exposure chambers via the diluter system (Brunson et al. 1998). Treatment solutions consisted of a control (base water with no Ni added) and a series of five 50% serial dilutions of Ni ranging from nominal concentrations of 31.25 to 500 μg/L Ni. A Ni stock solution was prepared by dissolving 23.98 g of nickel chloride hexahydrate salt in 3 L of 18.2 megaohm-centimeters water to result in a final concentration of 1,875 mg/L Ni; the stock solution was stored at room temperature throughout the test. A syringe pump was used to deliver 1 mL of this stock solution into 3,750 mL of base water (hardness 150 mg/L as CaCO3) with each cycle of the diluter system; this process created the greatest Ni test concentration (500 μg/L, termed High) for subsequent auto-dilution. The final Ni test concentrations were nominally 0 [Control], 31.25 [Low], 62.5 [Medium-low], 125 [Medium], 250 [Medium-high], and 500 [High] μg/L). The greatest nominal concentration (500 μg/L Ni) was selected to be representative of environmentally measured concentrations in mining-affected MCM waters (Allert et al. 2024). Subsequent concentrations were a product of preset dilutions based on the construction of the diluter system.

 Growth study

A 28-day chronic laboratory exposure was conducted to assess the effects of Ni on the growth of crayfish (Fig. 1b). At the initiation of the test, 96 crayfish of similar sizes were randomly selected, had wet weight taken, and were individually assigned to random exposure chambers within the diluter system (Fig. 1a). Mean wet weight (± one standard deviation) was 0.89 ± 0.17 g. Crayfish mortality was recorded daily for each exposure chamber. Excess food and waste were gently siphoned every 24 h from each chamber, but molts were left in the chambers to allow the crayfish the opportunity to consume calcium (Ca) and other nutrients to enable carapace hardening (Aiken and Waddy 1992). This was also done to reflect what would happen in a natural environment, since crayfish consume their molted exoskeletons in the wild. It is worth noting that this may have increased dietary consumption of Ni because Ni sorbs to the exoskeleton (Kouba et al. 2010). Daily and weekly water quality checks confirmed that uneaten food pellets left in the chambers for 24 hours had minimal effect on water quality. Full water quality methods and results are provided in Appendix A. Each exposure chamber received 0.05 g of sinking pellet food from test day 1 to day 25; animals were not fed on test days 26 and 27 (Fig. 1b). We chose 0.05 g as the starting weight of EP1 pellets daily for the growth study based on prior observations of crayfish being provided with different starting weights of EP1 pellets. This amount was chosen to ensure that each crayfish had an amount of food approximating what they could consume in 24 h. At the end of 28 days, all surviving crayfish were measured to determine the final wet weight and carapace length (Simon and Stewart 2014). Carapace length was measured with digital calipers (Tengyes IP54; Shenzhen Tengyes Technology Co. Ltd, Shenzhen, China) from the tip of the rostrum to the cervical groove at the end of the thorax (Hepper 1966). Carapace length was measured only at the end of the experiment to reduce handling stress imposed on the crayfish before initiation of exposures.

Concentrations of dissolved Ni, dissolved organic carbon (DOC), and major ions in the test waters were measured on test days 0, 14, and 28 in the Control, Low, Medium, and High exposure chambers; Medium-low and Medium-high concentrations were excluded from water quality analyses due to limited resources. A composite sample was produced by combining equal volumes of water from each of the four exposure chambers within a randomly selected aquarium per treatment in each diluter system. This sampling resulted in n = 8 water samples collected per sampling date. Waters were syringe filtered through 0.45 µm polyethersulfone membranes into appropriate pre-cleaned bottles and then preserved according to analysis type. The filterable fraction was selected because it is operationally defined as the dissolved, bioavailable fraction (USEPA 1993). Samples for Ni and major cation (sodium, Na; magnesium, Mg; potassium, K; Ca; manganese, Mn; Fe; strontium, Sr) analyses were aliquoted into low density polyethylene bottles and preserved by acidification with high purity nitric acid (2% volume/volume). Samples for major anion (fluoride, chloride, bromide, nitrite plus nitrate as nitrogen [N], phosphate, sulfate) analyses were stored in high density polyethylene bottles and preserved by refrigeration (4 °C); anion analyses were performed within 30 days of receipt of the sample. The DOC samples were aliquoted into amber glass vials and preserved by acidification (pH <2 with 9 N H2SO4) and refrigeration (4 °C); the maximum holding time for DOC was 28 days. Concentrations of Ni and major cations were measured using inductively coupled plasma-mass spectrometry; the method was established and validated under the USGS Quality Management System (QMS; USGS 2022) program and is similar to EPA Method 6020B (EPA 2014). Anion concentrations were quantified using anion exchange chromatography with suppressed conductivity detection (similar to EPA Method 9056A; EPA 2007) and DOC concentrations were measured as nonpurgeable organic carbon using combustion catalytic oxidation at 680 °C with nondispersive infrared detection (similar to EPA Method 415.3; Potter and Wimsatt 2005). The anion and DOC methods were developed and validated under the USGS QMS.

The experimental set-up for the nickel (Ni) exposure system. Exposures occurred in two flow-through diluter systems (Panel a), which included large, temperature-controlled water baths (labeled A). Twelve 75.7-liter (L) aquaria (labeled B) were placed into each water bath and connected to a diluter system. Within each aquarium, there were four 1-L glass exposure chambers (labeled C), which were covered with mesh screening to permit treatment solution exchange while preventing crayfish escape. Each exposure chamber contained a single crayfish, with n = 4 chambers per aquarium, and n = 48 crayfish within each diluter system, for n = 96 test animals. Each aquarium was equipped with opaque dividers (labeled D) between exposure chambers. Fresh treatment water was delivered hourly to each exposure chamber using a controlled delivery system (labeled E). A standpipe (labeled F) in each aquarium kept ambient water bath levels below the mesh screening of the exposure chambers. The study consisted of a chronic 28-day Ni exposure growth study (Panel b) followed immediately by consumption trials.

Alt Text: Diagram of the experimental set up and study timeline. Food consumption study

To determine whether different concentrations of waterborne Ni affected crayfish food consumption, a 24-h food consumption trial was conducted (Fig. 1b). Immediately following the 28-day chronic laboratory exposure growth study, crayfish were weighed and then placed back into the same exposure chambers for exposure to the same waterborne Ni concentrations. Note that the crayfish had been subjected to a 48-h fasting period, which began on test day 26 of the growth study (Fig. 1b), to ensure that all crayfish would be at similar levels of hunger, since 70% of crayfish intestinal contents are excreted in 12 h (McClain 2000). Animals were then fed 3.54 ± 0.02 g thawed, frozen bloodworms (Glycera sp.) and given 24 h to forage on the bloodworms. We selected 3.5 g as the starting weight of bloodworms for the food consumption study based on prior observations of crayfish being provided with different starting amounts of bloodworms. This amount was chosen to ensure each crayfish had more food available than they could consume in 24 h. Each crayfish was then weighed using a calibrated balance, measured for carapace length, and humanely euthanized via placement in a −20 °C freezer. Any remaining bloodworms were hand-picked from two to three randomly chosen exposure chambers within an aquarium and placed into weigh boats for wet weight determinations, as described in Appendix B. Crayfish excrement was not collected, and it was carefully distinguished from partially eaten bloodworms by color and consistency. Subsample weights were not taken from all four exposure chambers within each aquarium due to resource limitations; the total number of subsamples weighed was 69 out of N = 96 possible. Total consumption of bloodworms by each crayfish was calculated using the known starting weight of bloodworms, the weight of bloodworms removed from the exposure chamber at the end of the test, and an adjustment for water absorption by the worms between the start and end of the tests. Additional details regarding bloodworm weight adjustments and the tests used to derive those adjustments are presented in Appendix B.

 Data analyses

Crayfish wet weight change was calculated by subtracting initial wet weight from final wet weight. The level of significance was α = 0.05 for all tests, and all statistical analyses were performed in R (Version 4.4.0; R Core Team 2024). Normality was assessed using a Shapiro-Wilk test and visual inspection of quantile-quantile plots. Size results (initial weight, final weight, final length, weight change, and bloodworm consumption rate) were analyzed with a type III analysis of variance (ANOVA) using the function Anova() from R package ‘car’ (Fox and Weisberg 2019). Initial two-way analyses showed no effect of diluter system; therefore, diluter system was not included in the final analyses. A dose-response model was fit for wet weight change, final wet weight, and bloodworm consumption rate. The package ‘drc’ in R (version 3.0.1; Ritz et al. 2015) was used to estimate dose-response curve parameters using the function ‘drm.’ We used a four-parameter log-logistic model (eq. 1):

f (x) = c +d − c1 + eb(log(x) −log(e))

where x is the waterborne Ni concentration in microgram per liter, b defines the steepness of the curve, c is lower asymptote of the response curve, d is the upper asymptote of the response curve, and e is EC50 (the concentration at which 50% of organisms are affected) and log is the natural logarithm (Ritz et al. 2015). Root mean square error (RMSE) was calculated as a measure of fit for the dose response curves. Twenty percent effect concentrations (EC20s) were calculated in R with the function ‘drm’ (‘drc’ package; Ritz et al. 2015). In this study, we used change in crayfish wet weight over the 28-day chronic laboratory exposure to calculate the EC20 for juvenile growth. Raw data for all tests are available in Moore et al. (2025).

 Results Water chemistry

All water quality and major ion results were well within crayfish tolerance levels (Reynolds et al. 2013; Brown et al. 1995) and stable over the test duration (Appendix A); thus, only changes in waterborne Ni concentration might reasonably be expected to have affected growth and consumption rate results. Nickel concentration varied by treatment, but measured concentrations were close to nominal (Table 1). We measured 88.2%, 84.1%, 90.4% of the nominal concentration of Ni for the Low, Medium, and High treatments, respectively. The mean recovery (87.6%) was used to calculate an average conversion factor; we multiplied nominal Ni concentrations in all treatments by that conversion factor to obtain adjusted concentrations of Ni. This was required because Ni concentrations were not measured in all treatments. A post hoc analysis using the measured Ni concentrations for the Control, Low, Medium, and High treatments, and the adjusted nominal concentrations for the (unmeasured) Medium-low and Medium-high treatments demonstrated that the results (Appendix C) were not substantially different from the analysis using the adjusted nominal concentration for all treatments (Table 1). The difference between EC20s estimated by these two methods was <5%.

Treatment, test day, and nominal nickel (Ni) concentration (microgram per liter, µg/L) for crayfish (<italic>Faxonius virilis) </italic>test chambers over the 28-day chronic laboratory exposure for which Ni concentration was measured. Mean measured concentrations of waterborne nickel (Ni) were converted to adjusted nominal concentrations used for threshold estimations based on a mean Ni recovery of 87.6%. Concentrations are dissolved (0.45 micrometer filterable); means ± one standard deviation are for <italic>n </italic>= 2 replicates of each treatment.
Treatment Test day Nominal Ni (µg/L) Mean Measured Ni (µg/L) Adjusted nominal Ni (µg/L)
Control 0 0 0.24 ± 0.01 0
14 0 0.24 ± 0.04 0
28 0 0.27 ± 0.04 0
Low 0 31.25 27.5 ± 0.4 27.4
14 31.25 26.9 ± 0.5 27.4
28 31.25 28.3 ± 1.3 27.4
Medium-low NM 62.5 NM 54.8
Medium 0 125 103 ± 6 110
14 125 106 ± 6 110
28 125 107 ± 6 110
Medium-high NM 250 NM 219
High 0 500 452 ± 13 438
14 500 448 ± 11 438
28 500 457 ± 17 438

Note: NM indicates that this treatment was not measured for Ni concentration.

 Survival and growth

Survival was 99% across all treatments during the 28-day chronic laboratory exposure; only a single mortality occurred on test day 9 in the Medium-high treatment. There was no significant difference in initial crayfish wet weights across waterborne Ni concentrations (Table 2). However, growth—in the forms of wet weight change, final wet weight, and final carapace length—was significantly reduced with exposure to increased Ni concentrations (Table 2). Indeed, 80% or more of the variance associated with weight change and final weight were accounted for by Ni concentration (Table 2). Weight change decreased 17.8%, 51.6%, 68.0%, 67.7%, and 65.1.0% from control, for 31.25, 62.5, 125, 250, and 500 µg/L nominal exposure concentrations, respectively. The estimated EC20 for crayfish growth based on change in wet weight was 24.8 µg/L, and the estimated EC20 for crayfish growth based on final wet weight was 22.6 µg/L (Table 3). The 95% confidence interval for carapace length EC20 contained zero, indicating a lack of relationship between carapace length and EC20 (Table 3).

Stacked analysis of variance (ANOVA, Type III) results for five juvenile virile crayfish (<italic>Faxonius virilis)</italic> biological response variables, namely wet weight change, initial wet weight, final carapace length, final wet weight, and bloodworm consumption rate.
Response Term df SS F p-value R2
Wet weight change Intercept 1 3.816 245.118 <0.001 0.85
Ni concentration 5 1.626 20.896 <0.001 —
Residuals 18 0.280 — — —
Initial wet weight Intercept 1 3.237 608.587 <0.001 0.22
Ni concentration 5 0.027 1.028 0.431 —
Residuals 18 0.096 — — —
Final wet weight Intercept 1 14.081 602.035 <0.001 0.80
Ni concentration 5 1.676 14.327 <0.001 —
Residuals 18 0.421 — — —
Final carapace length Intercept 1 7208.6 9129.448 <0.001 0.55
Ni concentration 5 17.1 4.331 0.009 —
Residuals 18 14.2 — — —
Bloodwormconsumptionrate Intercept 1 117.756 2346.052 <0.001 0.82
Ni concentration 5 4.066 16.258 <0.001 —
Residuals 18 0.903 — — —

Note: Term is model term (nickel, Ni); df is degrees of freedom; SS is the sum of squares; F is the F-statistic; and R2 is the coefficient of determination.

Estimated 20% effect concentrations (EC20) in microgram per liter (μg/L) for juvenile virile crayfish (<italic>Faxonius virilis) </italic>based on adjusted nominal waterborne nickel (Ni) concentrations. Adjusted nominal concentrations were calculated using the mean Ni recovery of 86.7% (Table 1).
Response Predictor Estimate SE Lower 95% CI Upper 95% CI
Wet weight change Ni 24.812 4.922 14.544 35.079
Final wet weight Ni 22.641 6.616 8.839 36.442
Final carapace length Ni 31.584 15.754 -1.277 64.445
Bloodworm consumption rate Ni 46.105 15.780 13.190 79.021

Note: Response is the response variable in the dose response model. Predictor is adjusted nominal waterborne nickel concentration. The Estimate is the estimated concentration (μg/L) which would affect the response variable. The measures of precision for the Estimate are Standard Error (SE) and upper and lower 95% confidence intervals (CI).

 Food consumption rates

The consumption rate of bloodworms was inversely related to Ni concentration (Fig. 2). In other words, as the concentration of waterborne Ni increased, the rate at which crayfish fed on bloodworms decreased. Consumption rate decreased 5.8%, 8.8%, 25.9%, 32.8%, and 41.0% from control, for 31.25, 62.5, 125, 250, and 500 µg/L nominal exposure concentrations, respectively (Fig. 2). Concurrently, as final crayfish wet weight increased, consumption rates increased (Fig. 2). The estimated EC20 for impairment of bloodworm consumption rate by waterborne Ni was 46.1 µg/L Ni (Table 3).

During the 28-day chronic laboratory exposure growth experiment, crayfish were observed to have differing rates of consumption. Most crayfish in the control group were observed to eat 100% of their daily food allotment, whereas crayfish in the Medium and High treatments were often observed to have food left over the next day. However, the food pellets were designed to disintegrate in the water, so we were unable to accurately quantify the remaining food for that portion of this work.

Biological responses of juvenile virile crayfish (Faxonius virilis) to adjusted nickel (Ni) concentrations (87.6% of nominal concentrations; Table 1). Final crayfish wet weight was measured after 28 days of exposure to waterborne Ni. Consumption rates were measured over a 24-h period on test day 29, after crayfish had fasted for 48-h on test days 26–27. Panel a shows the final crayfish wet weight (grams [g]) modeled with a dose-response curve using the adjusted nominal waterborne Ni concentration (dissolved; microgram per liter [µg/L]; eq. 1). Nickel concentrations were natural logarithm (log)- transformed for analysis but are shown untransformed in the x-axis. The root mean square error (RMSE) was 0.182 and the EC20 was 22.6 µg/L. Panel b shows the relation between adjusted nominal Ni concentration (µg/L) and weight of bloodworms consumed (g) modeled by a dose response curve (RMSE = 0.135). As before, Ni concentrations were log-transformed for analysis but are shown untransformed in the x-axis; the EC20 for this relationship was 46.1 µg/L. Panel c shows the linear modeled relationship between consumption rate (g) and final wet weight (g); the correlation coefficient (R2) of this relationship was 0.67. Additional model parameter estimates are provided in (Table 2).

Alt text: a series of three line graphs showing the relationships between nickel concentrations and study endpoints. Discussion

The goal of this study was to determine whether chronic exposure to Ni at concentrations similar to those experienced by benthos in mining-affected ecosystems in southeast Missouri could adversely affect juvenile virile crayfish survival and growth. We examined the effects of waterborne Ni concentrations on survival and two metrics of growth (body weight and carapace length) in a laboratory setting over a 28-day chronic exposure period across all treatments, with only one death of N = 96 crayfish. This observation demonstrated that virile crayfish can survive concentrations of Ni ≥ 457 µg/L in moderately hard water (150 mg/L as CaCO3). This survival rate is supported by observations by Khan and Nugegoda (2007), who found that 327,000 µg/L Ni was needed to induce 50% mortality (LC50) in juvenile (4-week- old) common yabby crayfish (Cherax destructor) over an acute 96-hour exposure period. However, we have found that overall, the literature lacks evaluations of chronic exposure of juvenile crayfish to metals. Moreover, we found no other studies reporting lethal concentrations for chronic Ni exposure to crayfish; thus, the results of this study are critical for crayfish conservation and management. The United States hosts over 65% of the world’s crayfish species (Crandall and Grave 2017), so it remains crucial to evaluate how chronic exposure to contaminants—including metals—will impact crayfish survival, particularly because increased juvenile and adult mortality can reduce the success of wild populations (Meyer et al. 2007). Although Ni toxicity studies for crayfish are limited, studies for other benthic invertebrates have been published. For example, juvenile Fatmucket (Lampsilis siliquoidea; freshwater mussel) were found to have an EC50 for survival between 350 µg/L and 506 µg/L (hardness = 100 mg/L as CaCO3; Wang et al. 2017). Overall, our study suggests that Ni concentrations in the MCM are unlikely to cause virile crayfish mortality, but additional research could help to clarify effects of Ni on crayfish survival.

Unlike survival, we saw significant effects of Ni exposure on crayfish growth. Final wet weight, change in wet weight, and final length varied among Ni concentration treatments. However, usable EC20 estimates were only possible for wet weight change and final wet weight (EC20 =24.8 and 22.6 µg/L Ni, respectively); an EC20 for carapace length could not be reliably estimated. This was not unexpected; length may be impacted by the molt cycle status of an individual crayfish. During molting, length is non-continuous and occurs in a stepwise fashion while weight gain is more continuous (Chang et al. 2012), making it a more consistent endpoint. Over the course of the 28 days, Ni-exposed juvenile crayfish gained 17.8% to 68.0% less weight and were 10.7% to 38.8% lighter at the end of the trial compared to control crayfish (Fig. 2a). Such growth effects can alter ecological interactions between crayfish and other species. For example, when two species of crayfish are in competition, the faster-growing species typically has a greater survival rate (Kouba et al. 2021). Moreover, crayfish size can alter susceptibility to predation by fish and lead to increased anti-predator behaviors. For example, larger-sized virile crayfish (carapace lengths >30 mm), meant to be prey for predatory fish (Ambloplites, Perca, and Etheostoma spp.), spent more time moving and waving pincers in response to approaches by predators than crayfish with carapace lengths <30 mm (Keller and Moore 2000). Fecundity and egg size increase with increasing female crayfish body size, potentially impacting recruitment rates (Nakata and Goshima, 2004; Maguire et al. 2005). Therefore, the reduced growth demonstrated by our study has the potential to lead to adverse effects on populations and communities.

One potential factor for the observed reduced growth was the reduced consumption rates. We found that food consumption rates decreased with increasing concentrations of dissolved Ni (Fig. 2b; Table 2). Crayfish exposed to the greatest Ni concentration (adjusted nominal Ni, 438 µg/L; Table 1) consumed 41% less food than the controls. While these findings were significant, it is important to note that our study was limited by a small sample size (n = 4 aquaria with 2-3 subsamples per aquarium) and a short time frame (24 h). A consumption study occurring concurrently with the growth study would have been optimal; however, as previously mentioned, the pelleted food we selected for the growth study disintegrated in water. Therefore, the consumption study had to take place after the growth study with a more structurally-sound food: bloodworms. Future crayfish consumption studies might usefully consider testing several types of food sources, because we hypothesize that consumption rates could vary by food source.

Altered feeding behaviors of crayfish in the presence of metals has been described in other studies. For example, the behavioral response of red swamp crayfish (Procambarus clarkii) and rusty crayfish (F. rusticus) to a feeding stimulant was suppressed during a five-minute exposure to a mixture consisting of Cu, chromium (Cr), As, and selenium (Se) at a total metals concentration of 98 µg/L compared to control animals (Steele et al. 1992). In the presence of 20 µg/L Cu, Appalachian brook crayfish (C. bartonii) did not seek out food or required significantly longer time to find food in a Y-maze setting compared to crayfish not exposed to Cu; it was hypothesized that crayfish exposed to Cu lost the ability to detect food, and delayed effects were observed after removal of the toxicant (Sherba et al. 2000). Sublethal levels of Cu exposure have been shown to result in impaired chemoreception and orientation, as well as slower movements, in rusty crayfish (Lahman et al. 2015). Our findings of reduced consumption rates in Ni-exposed virile crayfish were consistent with these findings for other metals, but we did not determine a mechanism.

The relationship between consumption rates and growth is supported by our finding that Ni concentration and final crayfish wet weight were both good predictors of consumption rates, but it is possible that other physiological impacts of Ni exposure contributed to reduced growth. For example, sublethal metals exposures have been found to inhibit the central nervous system (Devi et al. 1996; Danilovic et al. 2022) and influence circadian rhythmicity of heart rates of crayfish (Styrishave et al. 1995). Furthermore, sublethal concentrations of Pb and Cd impaired oxidative enzymes in the digestive gland and gills of noble crayfish (Meyer et al. 1991). Thus, more research is needed to understand the physiological impacts and potential mechanisms of Ni toxicity and the mechanisms of reduced growth.

In addition, there are other endpoints which may affect or be affected by Ni toxicity, such as molting and reproduction. Crayfish molting—which is sensitive to many factors, including temperature, nutrition, light, stress, and reproductive state (Gutiérrez-Yurrita and Montes 1999; Aiken and Waddy, 1992)—can lead to increased sensitivity to metal exposure (Wiggington and Birge, 2006). Molting was observed in crayfish throughout the study but was not tracked, so it was not possible to determine whether Ni in our experiments affected molting or whether molting status affected toxicity. Reproduction may be especially sensitive to metals toxicity as a result of its high metabolic cost (Gashkina 2024). At a concentration of 17,300 µg/L (pH = 7.4), Ni exposure disrupted oogenesis in Danube crayfish ovaries, ultimately altering and disrupting reproductive organs and cellular processes (Zarnescu et al. 2017). Previous studies of chronic exposure of aquatic invertebrates to waterborne Ni have indicated that reproduction is typically the most sensitive endpoint, while biomass, survival, and growth are less sensitive (Zarnescu et al. 2017; Wang et al. 2020). Research into the effects of Ni on virile crayfish and other species would help to further elucidate the effects of toxicity on individuals and ecosystems.

As a tool for helping to guide resource management decisions, this study establishes methods to evaluate baseline growth and consumption rates of virile crayfish and other crayfish species in a laboratory setting. It also documents those characteristics for juvenile virile crayfish of the 0–1 year age class. The data collected by this study could be used to provide a foundation for future bioenergetics modeling and incorporation of metals contamination into models. An example application of bioenergetics modeling with crayfish is the dynamic energy budget modeling of juvenile red swamp crayfish fed four different diets (Gutiérrez-Yurrita and Montes 1999). Bioenergetic modeling allows for extrapolation of results from laboratory-based toxicity testing across the lifespan of an individual and across a population (Wildhaber et al. 2017a, b). It can be a useful tool to understand the effects of management actions on a population; further, it can be used to predict future rates of invasive crayfish dispersal and survival of native crayfish populations. This type of work has been done on fishes in the Missouri River (Wildhaber et al. 2025, Wildhaber et al. 2024) and would be valuable for crayfish worldwide.

From a community ecology perspective, species that can chronically tolerate elevated concentrations of Ni, or other metals and contaminant types could have a competitive advantage. Interspecies conflict between crayfish is often intense, and dominant species will outcompete nondominant species for optimal habitat (Garvey et al. 1994). Virile crayfish were previously found to be the most tolerant to Cd exposure of six tested crayfish species (Wigginton and Birge 2007). However, our study is the first to test virile crayfish with Ni. If it turns out that virile crayfish are more tolerant of Ni and potentially other metals than other species, virile crayfish will likely have an advantage when competing with both native and invasive crayfish species having lower tolerances in contaminated areas.

Effects of Ni on crayfish may translate to indirect effects on aquatic communities. Reduced consumption rates are particularly notable because this effect could drive ecological interactions of native and invasive crayfish. Juvenile crayfish generally filter-feed on microscopic planktonic organisms, transitioning to larger food sources like macrophytes and macroinvertebrates as they grow (Velez-Gavilan 2016). Additionally, juveniles are typically more sensitive to metal toxicity than adults (Knowlton et al. 1983; Eversole and Seller 1997; Wigginton and Birge 2007). This relationship between growth and ecological niche could lead to altered food web dynamics, productivity, and ecosystem services, particularly if crayfish were to reduce food consumption rates or utilize different prey in Ni-contaminated areas. Any shift in crayfish behavior is likely to influence multiple trophic levels and community-level properties (Danilovic et al. 2022). Further, crayfish are often consumed by fish, birds, turtles, and snakes (NEMESIS 2023), so Ni uptake in crayfish tissues could potentially propagate into the upper trophic levels of the aquatic food web (Danovaro et al. 2023) and be transferred into terrestrial ecosystems. It will be critical to determine lethal and nonlethal responses, including toxicity thresholds, of various crayfish and other aquatic species to both single metals and metals mixtures. Such research could help evaluate the potential effects of competitive interactions among aquatic organisms in contaminated environments. Ultimately, understanding the effects of metals contamination on keystone taxa like crayfish could assist natural resource managers with implementing data-informed best practices to preserve the ecosystem services of native species and prevent invasions of nonnative species.

 Acknowledgements

We thank all who helped from the U.S. Geological Survey (USGS), particularly Vanessa Melton for assistance with major anion and dissolved organic carbon analyses; Ben West and Nicole Tripp for editorial suggestions that helped improve the manuscript; and Ben Bates and Jack May for assistance with the study. We also thank David Soucek (USGS) and the anonymous journal peer reviewers for their helpful comments. Any use of trade, firm, or product names is for descriptive purposes only and does not imply endorsement by the U.S. Government.

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During the 28-day chronic laboratory exposure, dissolved oxygen and temperature (YSI Pro-ODO meter with Optical DO probe; YSI Incorporated, Yellow Springs, Ohio, USA), pH (HACH HQ40d meter with a PHC201 probe; HACH Company, Loveland, Colorado, USA), and conductivity (HACH HQ40d meter with a CDC401 probe; HACH Company, Loveland, Colorado, USA) were measured daily in a random exposure chamber in each aquarium according to standard methods (ASTM International 2000a).

For each of the four weeks of the study, composite samples of water were used to measure ammonia, hardness, and alkalinity. The first step of creating composite samples was randomly selecting four aquaria each week from each of the two diluters, with one aquarium per diluter representing each of the treatment levels: Control, Low, Medium, and High (eight aquaria per week total). Approximately equal volumes of water from each 1-L exposure chamber in a single aquarium were combined to make a composite sample. Ammonia was measured by colorimetry (Eaton et al. 2005; Hanna Iris Visible Spectrophotometer HI801, Hanna Instruments, Woonsocket, Rhode Island, USA). Alkalinity and hardness were quantified using titrimetry (ASTM International 2000b).

Test water quality characteristics were generally consistent across the 28-day chronic laboratory exposure and consumption rate test phases, with the exception of dissolved oxygen (DO) concentrations in the consumption test (Moore et al. 2025). The mean water temperature (± one standard deviation) was 21.1 ± 0.2 °C (n = 720). Mean water quality measurements were pH = 7.96 ± 0.20 (n = 720), specific conductance = 325 ± 18 µS/cm at 25 °C (n = 720), hardness = 150 ± 1 mg/L as CaCO3 (n = 38), alkalinity = 126 ± 3 mg/L as CaCO3 (n = 40), and total ammonia nitrogen = 0.063 ± 0.034 mg/L (n = 40). Mean DO concentrations were 8.23 ± 0.27 mg/L during the 28-day chronic laboratory exposure but were consistently lower across treatments at 5.88 ± 0.79 mg/L at the end of 24-h consumption rate test. However, virile crayfish can survive DO levels as low as 1.5 mg/L (Brown et al. 1995), so we did not expect reduced DO to have significantly altered our consumption results. Mean concentrations for major ions in the test waters (n = 24) were Ca = 34 ± 1 mg/L, chloride = 16 ± 1 mg/L, K = 1.3 ± 0.1 mg/L, Mg = 13 ± 1 mg/L, Na = 12 ± 1 mg/L, sulfate = 28 ± 1 mg/L, and Sr = 0.18 ± 0.01 mg/L. Concentrations of bromide, DOC, Fe, Mn, nitrite plus nitrate as N, and phosphate were below method reporting limits or method limits of identification (Moore et al. 2025). Fluoride concentrations were at or near the 0.2 mg/L reporting limit for all samples.

 Bloodworm tests and adjustment equations Methods for bloodworm tests

Determination of the ending weight of bloodworms during the food consumption study was not straightforward and ultimately required extrapolation. There were two separate issues regarding bloodworms: a change in water removal methods and water absorption by the worms. To account for an inadvertent change in bloodworm weighing methods between the start and end of the food consumption study, we compared two methods of water removal when weighing bloodworms. The methods were termed ‘no wicking’ and ‘wicking.’ The ‘no wicking’ approach involved letting gravity drain water away from each pile of bloodworms by tilting a weigh boat at a slight angle until it appeared that water stopped draining. We used this method applied to thawed bloodworms before they were added to the exposure chambers at the start of the consumption study. ‘Wicking’ involved taking absorbent laboratory wipes and slightly touching the bottom of the bloodworm pile to absorb more water than would drain from the bloodworms by gravity. For the first phase (comparison of water removal methods), we weighed 16 similar sets of thawed bloodworms (3.51 ± 0.01 g; mean ± 1 SD). Treatments were base water (150 mg/L hardness [as CaCO3] with no Ni added) and water with an adjusted nominal Ni concentration of 438 µg/L (n = 8 bloodworm sets per treatment). For the second phase (water absorption by bloodworms), we weighed 16 sets of thawed bloodworms in four different weight groups: 0.11 ± 0.0 g, 1.22 ± 0.01 g, 2.31 ± 0.01 g, or 3.51 ± 0.01 g (n = 4 bloodworm sets for each weight group). Two sets per weight group were added to each of the base water and 438 µg/L Ni treatments. Results of the water removal and absorption trials were used to generate linear equations intended to convert the ‘wicking’ final bloodworm weights from the consumption study to values equivalent to ‘no wicking’ initial weights.

 Bloodworm test equations and applications

Regarding water removal, the linear regression equation of ‘no wicking’ final wet weight (y) and ‘wicking’ final wet weight (x) was y = 1.085x − 0.025 (Equation B1, r2 =0.997, n = 32). This regression equation provided the basis to use the ‘wicking’ final weights to approximate ‘no wicking’ final weights. For water absorption, the regression equation of ‘no wicking’ initial wet weight (z) and ‘no wicking’ final wet weight and was z = 0.776y + 0.073 (Equation B2, r2 = 0.976, n = 32). This regression equation was the basis for approximation of the equivalent of initial weights from ‘no wicking’ final weights. These equations were then applied in sequence (Equation B1 followed by Equation B2) to the final weights of bloodworms after the consumption study to convert final wet weights to a value comparable to initial wet weights.

 Analysis using alternative Ni concentrations

In the Results section of the main text, we calculated EC20 values using adjusted nominal Ni concentrations for all treatments, based on the average Ni recovery of 87.6%. In the following analysis, we calculate EC20s using the measured Ni concentrations for the Control, Low, Medium, and High treatment levels, and applied the nominal adjustment to only the Medium-low and Medium-high treatments, which were not directly measured due to limited resources (Table C1). Comparisons of the EC20s estimated using all adjusted nominal concentrations (Table 3) and EC20s using a mixture of measured and adjusted Ni concentrations (Table C2) demonstrate that the differences between the results were negligible.

Treatment, test day, and nominal Nickel (Ni) concentration (conc.; microgram per liter) for virile crayfish <italic>(Faxonius virilis) </italic>test chambers over the 28-day chronic laboratory exposure for which Ni concentration was measured. Mean measured concentrations of waterborne Ni (denoted as M) were converted to percent recovery to obtain an adjustment factor (87.6%) to derive the adjusted nominal concentration for unmeasured treatments (denoted AN).
Treatment Test day Nominal Ni conc. Mean measured Ni conc. Percent recovery of measured Ni Measured (M) or adjusted nominal (AN) Ni conc.
Control 0 0 0.24 ± 0.01 — 0.25 (M)
Control 14 0 0.24 ± 0.04 — —
Control 28 0 0.27 ± 0.04 — —
Low 0 31.25 27.5 ± 0.4 88.2 27.6 (M)
Low 14 31.25 26.9 ± 0.5 — —
Low 28 31.25 28.3 ± 1.3 — —
Medium-low NM 62.5 NM — 54.8 (AN)
Medium 0 125 103 ± 6 84.0 105 (M)
Medium 14 125 106 ± 6 — —
Medium 28 125 107 ± 6 — —
Medium-high NM 250 NM — 219 (AN)
High 0 500 452 ± 13 90.4 452 (M)
High 14 500 448 ± 11 — —
High 28 500 457 ± 17 — —

Note: Measured concentrations were dissolved (0.45 micrometer filterable); means ± one standard deviation are for n = 2 replicates of each treatment. NM indicates that this treatment was not measured for Ni concentration; — indicates that a percent recovery was not calculated because Ni was not measured in this treatment.

The estimated 20% effects concentrations (EC20) in microgram per liter (μg/L) for juvenile virile crayfish (<italic>Faxonius virilis) </italic>based on the use of measured (Control, Low, Medium, and High treatments) with adjusted nominal waterborne (Medium-low, Medium-high) nickel concentrations. Response is the response variable in the dose response model and estimate is the EC20 for each variable. The measures of precision for the Estimate are Standard Error (SE) and upper and lower 95% confidence intervals (CI). Percent difference indicates the percent difference from estimates in Table 3 for using all adjusted nominal concentrations in the EC20 estimate.
Response Estimate SE Lower 95% CI Upper 95% CI Percent Difference
Wet weight change 25.053 4.902 14.828 35.278 0.97
Final wet weight 22.858 6.565 9.164 36.552 0.96
Final carapace length 30.194 13.486 2.062 58.326 -4.40
Bloodworm consumption rate 45.150 15.426 12.972 77.329 -2.07
Additional Information

Author Contribution Statement: Author Contributions: Mark L. Wildhaber and Adrian P. Moore contributed to study conception and design. Adrian P. Moore, Mark L. Wildhaber, Zachary D. Beaman, Kendell R. Bennett, Karlie K. Ditter, and Danielle Cleveland contributed to material preparation and data collection. Statistical analyses were performed by Adrian P. Moore, Mark L. Wildhaber, and Tyler J. Grant; Danielle Cleveland quantified nickel and major cations in the water samples. The first draft of the manuscript was written by all authors and all authors commented on subsequent versions of the manuscript. All authors read and approved the final manuscript.

Data Availability: All raw data used in the development of this manuscript are published in Moore et al. (2025), accessible at https://doi.org/10.5066/P13X7LX7.

Funding: This work was supported by the U.S. Geological Survey Biological Threats & Invasive Species Program and Land Management Research Program.

Ethics approval: All animal laboratory testing protocols were approved by the USGS Columbia Environmental Research Center Animal Care and Use Committee and conformed to AFS/AIFRB/ASIH (2014) Guidelines for the Use of Fishes in Research.

Conflicts of Interest: The authors declare no conflicts of interest.