Effects of Temperature, Total Dissolved Solids, and Total Suspended Solids on Survival and Development Rate of Larval Arkansas River Shiner

The rivers and streams flowing through the Great Plains of central North America that once hosted high levels of biodiversity are now considered among the most imperiled biomes on the continent (Dodds et al. 2004). The native ichthyofauna of the Great Plains has experienced high levels of local extirpations influenced by changes in the physicochemical environment (Hoagstrom et al. 2011). For example, average temperature within the Great Plains ecoregion has increased by approximately 0.8°C relative to the 1970s and is predicted to continue to increase by another 1.0–7.5°C by the end of the century (Fields et al. 2007; Karl et al. 2009). Increases in air and water temperatures interact with anthropogenic water withdrawals to intensify stream drying and concentrate total dissolved solids (TDS; Hammer 1971; Williamson et al. 1999; Dodds et al. 2004). Total suspended solids (TSS) within Great Plains will continue to increase as a result of soil erosion after intense rainfall events (Leemans and Kleidon 2002). Although the changes projected for Great Plains rivers do not seem to present an improved outlook for already imperiled fishes, there is little information available to understand how species might respond to these predicted changes.

The plight of pelagic broadcast-spawning, or pelagophilic, cyprinids inhabiting the rivers of the Great Plains ecoregion represents a nexus where anthropogenic modifications, including climate change, may affect their early life-history stages and ultimately determine their persistence. Pelagophilic cyprinids are a reproductive guild of approximately 20 species of small-bodied (<7-cm) fishes that produce semibuoyant, nonadhesive embryos (Moore 1944; Bestgen et al. 1989; Platania and Altenbach 1998; Bonner and Wilde 2000; Perkin and Gido 2011). Although multiple factors have been proposed as contributing to unsuccessful reproduction in pelagic broadcast-spawning cyprinids, much still remains unknown about how projected physicochemical changes in temperature, TDS, and TSS, might affect development and survival of their early life-history stages.

Arkansas River Shiner Notropis girardi was listed as a federally threatened species in 1998 pursuant to the U.S. Endangered Species Act (ESA 1973, as amended; USFWS 1998). Although they are among the better studied of the pelagophilic cyprinids, uncertainties regarding their ecological requirements and response to changing environmental conditions limit conservation and recovery planning (Wilde 2002). This species is endemic to the Arkansas River basin but has been extirpated from much of its historical range over the past 30 y (Worthington et al. 2014). Although numerous factors have been proposed to explain this rapid range-wide decline, there is still a need to better understand how the changing physicochemical environment may also be contributing. For example, reductions in stream flow have negative effects on reproductive success of Arkansas River Shiners (Durham and Wilde 2009), but the mechanisms responsible for decreasing reproductive success remain unclear. In addition to influencing transport times and retention, reductions in stream flow could produce changes in physicochemical conditions unrelated to current velocity that may affect reproductive success of pelagophilic cyprinids. Numerous studies have demonstrated the effects that physicochemical conditions, such as temperature, TDS, and TSS have on the abundance and persistence of Arkansas River Shiner and other pelagophilic cyprinids (Matthews and Hill 1980; Polivka 1999; Ostrand and Wilde 2002, 2004), but the effects on their developmental rate and larval survival are not well-understood. A thorough understanding of how Arkansas River Shiners respond to temperature, TDS, and TSS during their early life history is an important component necessary for developing an effective conservation program that is responsive to the dynamic environment of the Great Plains. Therefore, the objectives for our study were to 1) assess the effects of temperature, TDS, and TSS on the developmental and survival rates of Arkansas River Shiner embryos and larvae; 2) create an early life-history stages identification guide for Arkansas River Shiner.

We captured adult Arkansas River Shiner in June 2012 using seines from 1) the South Canadian River at the US-283 crossing near Roll, Oklahoma (n = 54; personnel from U.S. Fish and Wildlife Service); and, 2) the Pecos River approximately 10 km upstream from the US-70 crossing in Chaves County, New Mexico (n = 56; personnel from Museum of Southwestern Biology). We transported all individuals (n = 110) by truck using small aerated tanks to the U.S. Fish and Wildlife Service Tishomingo National Fish Hatchery in Tishomingo, Oklahoma. We housed the broodstock in a shaded flow-through raceway system maintained at ambient water temperatures (19.9°C–27.9°C) and photoperiod cycle of 15 h light and 9 h dark. Hatchery personnel fed the Arkansas River Shiner freeze-dried Tubifex and frozen chironomid larva (bloodworms) daily.

We induced spawning following an established protocol for captive breeding Arkansas River Shiners (K. Graves, Hatchery Manager, U.S. Fish and Wildlife Service, personal communication). Briefly, we anesthetized the fish by immersing them into a 25-mg/L tricaine methanosulfate (Western Chemical, Inc., Ferndale, WA) solution. To determine sex, we applied pressure to the abdomen of each fish sufficient to express gametes then separated individuals by sex into two 37.9-L aquaria (Aqueon Products, Franklin, WI) maintained at ambient temperature. We allowed the fish to recover undisturbed for 36 h before inducing them to spawn.

On 26 July 2012 at approximately 0830 hours, we induced spawning in 10 adult Arkansas River Shiners of each gender. To induce spawning, we anesthetized fish and after loss of equilibrium (1–2 min), we then transferred each individual to a wet gauze-pad cradle on the stage plate of a dissection microscope. We injected approximately 1.0 mL of a 100-mg/L solution of ground carp pituitary extract (Argent Laboratories, Redmond, WA) dissolved in sterile saline along the ventral midline near the anus of each fish. After injection, we held all fish in a 10.8-L plastic tub until they regained equilibrium. Then, we transferred the recovered adults to a 37.9-L aquarium and left them undisturbed to allow reproduction to occur. As a result, all embryos from all adult pairings were mixed together.

Once spawning was finished and the perivitelline space began to fill, the embryos became semibuoyant and were ready for transfer to the eight temperature–TDS–TSS treatment beakers. For each treatment, we collected embryos from the spawning tanks using a clear polyvinyl chloride pipe with a 12.7-mm diameter attached to a clear flexible polyurethane tube of the same diameter, and placed them into a full 100-mL graduated cylinder. Once the embryos displaced 1.0 mL of water from the graduated cylinder, we transferred the contents to a 240-mL glass treatment beaker using a 3-mL plastic pipette. Preliminary work determined that approximately 45 Arkansas River Shiner embryos occupied a volume of 1.0 mL (Mueller 2013). We therefore estimated that 45 embryos were exposed to each treatment.

We chose two treatment levels of temperature (25°C, 31°C), TDS (1,000 mg/L, 6,000 mg/L), and TSS (0 mg/L, 3,000 mg/L) to represent the expected range of typical and projected environmental conditions that Arkansas River Shiner in the Canadian River experience during their spawning season. We selected temperature treatment levels to encompass a range in which successful spawning was observed in laboratory settings (Platania and Altenbach 1998; K. Graves, U.S. Fish and Wildlife Service, personal communication) and were within the range of mean monthly water temperatures observed in the Canadian River during Arkansas River Shiner spawning season (April 2011–July 2012, Gage Number 07227500; USGS 2014). We selected the high treatment levels for TDS and TSS to represent high values observed in the Canadian River (TDS = 6,450 mg/L, TSS = 3,033 mg/L; Pigg et al. 1999; Gage Number 07227500; U.S. Geological Survey). We used the 25°C-1,000 mg/L TDS–0 mg/L TSS level as the control for the experiment because it represented the same conditions as the spawning tanks.

We created the temperature treatments by adding a 50-W submersible heater and thermostat (Aqueon Products) to the bottom of two 37.9-L aquaria that were filled with approximately 6.5 L of water. We placed the heater horizontally on the backside of the tank and we filled the tanks with water to just below the tops of the beakers so no additional water would get into our treatments. We placed a single 304.8-mm-long air diffuser (Top Fin 12-Inch Aquarium Air Stone, PetSmart, Phoenix, AZ) horizontally in each aquarium in front of the heater to help circulate water and ensure that temperature was uniform throughout the tank. Using a portable fish tank thermometer (Top Fin Digital Aquarium Thermometer, PetSmart), we checked the temperature in the tank randomly throughout the study. In both aquaria, at the opposite end of the heater we placed four 250-mL glass beakers. These beakers housed the TDS–TSS treatment groups. We checked the temperature within each beaker frequently during the study using our portable fish tank thermometer. The high TDS level was established through the addition of Instant Ocean seawater mix (Spectrum Brands, Cincinnati, OH). We used commercially packaged Moroccan red clay (i.e., rhassoul) to obtain the desired TSS levels as described by Zamor and Grossman (2007) and Hazelton and Grossman (2009). We weighed both the seawater mix and red clay to the nearest 0.01 g using a CQT core compact portable electronic balance (Adam Equipment, Inc., Danbury, CT) and added both to a premeasured volume of water. To ensure the red clay and Arkansas River Shiner embryos remained in suspension for the duration of the study, we placed a 25.4-mm air diffuser (Grreat Choice Mini Air Stone, PetSmart) at the bottom of all eight beakers. In addition to the air diffuser, we used a glass stir rod to gently stir the water in each beaker once per hour to resuspend anything that may have settled to the bottom.

We collected embryos from each beaker at 1, 2, 4, 8, 12, 16, 24, and 36 h postfertilization using a plastic pipette and placed them into 1.5-mL plastic, clear, snap-cap containers containing a 3% formalin solution. We labeled all containers with the appropriate temperature–TDS–TSS treatment and held in ziploc bags separating them by hours postfertilization. At the 36-h mark, we drained all beakers into a fine mesh net to ensure no living larvae were left in our treatment beakers. We added any remaining larvae to our 36-h postfertilization results. We used a fine mesh net with a mesh size of 0.09 cm to minimize abrasion while collecting the remaining larvae. During each collection hour, when we collected >5 embryos in a pipette, we retained them all to ensure that potentially damaged embryos were not returned to the treatment beakers. This resulted in inconsistent sample sizes for each sampling period (Table S1, Supplemental Material). Furthermore, we were unable to acquire a sufficient number of embryos for a fully replicated factorial design from induced spawning. Therefore, Arkansas River Shiner developmental data are represented by a single, unreplicated experiment.

At the conclusion of our study, we transferred the plastic bags holding all Arkansas River Shiner embryos to Texas Tech University (Lubbock, TX) for further analysis of developmental stage. We used a dissecting microscope (SZX16, Olympus Corporation, Tokyo, Japan) equipped with a digital camera (Infinity 1, Lumenera Corporation, Ottawa, ON, Canada) to capture images of all preserved specimens under 11.5× magnification. To aid in the determination of developmental stages, we enhanced images by using the unsharp mask filter and the brightness adjustment feature of the ImageJ v.1.46 software (Abràmoff et al. 2004).

We based assignments of developmental stages primarily on the criteria presented by Moore (1944) for Arkansas River Shiner and supplemented with descriptions of other species in the Cyprinidae family (i.e., Spottail Shiner Notropis hudsonius, Jones et al. 1978; Speckled Chub Macrhybopsis aestivalis, Bottrell et al. 1964; Rosyface Shiner Notropis rubellus, Reed 1958; Zebrafish Danio rerio, Kimmel et al. 1995; Grass Carp Ctenopharyngodon idella, Black Carp Mylopharyngodon piceus, Silver Carp Hypophthalmichthys molitrix, and Bighead Carp Hypophthalmichthys nobilis, Yi et al. 1988; Chapman 2007). Although some of these species have different reproductive strategies compared to that of the Arkansas River Shiner, we expected that the stages of early development would be consistent among cyprinids.

We classified images (n = 227) of Arkansas River Shiner embryos (prehatching developmental stages) and larvae (posthatching developmental stages) into 14 distinct developmental stages (Table 1; Figure 1). Hatching occurred between the caudal fin appearance stage and the otolith appearance stage (Figure 1). When analyzing our developmental rate data, we combined stages that lacked an adequate number of replicates with others, dropping our total number of stages to nine (Figure 2; Table S1, Supplemental Material). To obtain larger sample sizes, we combined early, mid, and late gastrula together as gastrula stage; tail bud and caudal fin together as tail bud stage; and otolith appearance, melanoid eye, and gas bladder emergence together as otolith appearance stage. Early development of Arkansas River Shiner was rapid, with most individuals entering the late gastrula or neurula stages within 4 h postfertilization and hatching within 24–36 h. Embryos from the 31°C temperature treatment tended to be more advanced in their development at a given time than their counterparts incubating at 25°C (Figure 2). Individuals in the highest temperature–TDS–TSS treatment group began hatching as early as 8 h postfertilization (Figure 2). Embryos developing at higher TDS and TSS treatment levels tended to reach developmental stages earlier than those at lower TDS–TSS treatment levels in the same temperature (Figure 2). Furthermore, the total number of embryos and larvae classified into developmental stages from the lower temperature treatment levels (n = 144) was almost double those from the higher temperature treatment levels (n = 83) because of a lack of individuals at the later sampling periods. These results suggest that Arkansas River Shiners raised in the 25°C treatments experienced a mortality rate of approximately 20% compared with approximately 54% for the individuals exposed to the 31°C treatments.

Our study represents an initial step in the process of determining how the changing physicochemical environment of the rivers of the Great Plains ecoregion may be affecting the development and survivability of Arkansas River Shiners. With the climate and landscape use changes being made to the Great Plains we can expect to experience increases in temperature, TDS, and TSS within our river systems. The presence and abundance of pelagophilic cyprinids, such as Arkansas River Shiner, exhibit strong correlations with these physicochemical factors. For example, Ostrand and Wilde (2002) found that plains minnow Hybognathus placitus, smalleye Shiner Notropis buccula, and sharpnose Shiner Notropis oxyrhynchus relative abundances were greatest at sites in the upper Brazos River in Texas with low TDS and high TSS. Furthermore, the distribution of Arkansas River Shiner in the South Canadian River in Oklahoma was negatively correlated with both temperature and TDS (Polivka 1999). Our results suggest that changes in the temperature, TDS, and TSS in the rivers and streams of the Great Plains ecoregion may affect the persistence of these species by influencing the developmental rate and survival of early life-history stages.

Even though we were unable to produce a sufficient number of embryos for a fully replicated factorial experiment, the protocols used to spawn Arkansas River Shiner for this study did produce viable embryos. We experienced difficulties collecting a species listed as threatened, which resulted in delayed collections and a smaller than expected sample size. Additionally, previous spawning attempts by these fish may have reduced the number of gametes each individual could produce. Future work should induce as many individuals of Arkansas River Shiner as possible at the beginning of their spawning season. This would help increase the number of viable embryos and also allow for multiple spawning events to occur during their spawning season if needed. As our study came to a close, meeting our goal of collecting five embryos in each exposure group became more difficult to achieve for those embryos incubated at 31°C. Although we used the same volumetric method as described in our methods for placing embryos into each beaker, this technique had its limitations and did not ensure that all beakers would start with the exact same number of embryos. These results may suggest that although higher temperatures result in faster development, which may be beneficial in shorter river segments, this faster development seemed to also be associated with higher levels of mortality. Though our study did not directly assess mortality rates at regular intervals in each treatment, the number of embryos and larvae recovered for classification at the end of treatment allowed an estimate of the mortality rates associated with different temperatures.

Although a strong relationship between temperature and developmental rate was expected, the increased developmental rate associated with elevated TDS and TSS treatments was not anticipated. However, a similar relationship between TDS and developmental rate has been described in the Rio Grande silvery minnow Hybognathus amarus (Cowley et al. 2005). When incubating in high levels of TDS, Rio Grande silvery minnows hatched hours earlier than individuals at the same temperature and lower TDS levels. However, the mechanism(s) driving this relationship has not yet been determined. Embryos of another cyprinid, zebrafish, hatched earlier when exposed to elevated TDS levels produced by inorganic limestone and limestone in suspension than control groups (Reis 1969).

Sorensen et al. (1977) found that increases in suspended solid loads tend to coincide with decreases in dissolved oxygen, which could lead to hypoxic stress during development in many fish species. However, the timing of exposure during early development to increases in TSS may contribute to how the species are affected. For example, Muncy (1979) found that embryos of warm-water fishes show no effects of exposure to high levels of TSS during water hardening of the eggs, but these same levels could result in mortalities when oxygen demands of the embryos were greater during later stages of development. Siefert et al. (1974) found that Smallmouth Bass Micropterus dolomieu development was accelerated by low dissolved oxygen concentrations associated with increased TSS. In salmonids, fry survival can decrease as much as 3.4% for each 1% increase in fine sediment (Cederholm et al. 1981). Although rapid development for Arkansas River Shiner and other pelagophilic cyprinids could be a strategy of survival (Platania and Altenbach 1998), the oxygen demands associated with the higher metabolic rates that are likely necessary to support rapid development may render these species more susceptible to mortality or other negative effects from elevated levels of TDS and TSS.

Although Arkansas River Shiner embryos reached developmental milestones faster in higher temperatures, future work addressing the survival of their early life-history stages under a range of conditions is warranted. Under the projected changes due to global climate change, a better understanding of how they are affected by the ever-changing environment of the Great Plains, including changes in temperature, TDS, and TSS, is needed. The distribution of adult Arkansas River Shiners seem to be negatively correlated with TDS and TSS (Matthews and Hill 1980; Reash and Pigg 1990; Polivka 1999), and our results suggest that this may be in part due to high mortality rates in their embryo and larvae stages. A better understanding of how shifting physicochemical conditions, such as temperature, TDS, and TSS affect the early life-history stages of different species is important. Such an understanding of different species will allow us to better predict stream reaches capable of supporting these species even as water quality in the rivers and streams of the Great Plains continue to change.

Please note: The Journal of Fish and Wildlife Management is not responsible for the content or functionality of any supplemental material. Queries should be directed to the corresponding author for the article.

Table S1. Developmental stage of Arkansas River Shiner Notropis girardi, eggs and larvae, incubating in different temperature (25°C, 31°C), total dissolved solid (TDS; 1,000 mg/L, 6,000 mg/L), and total suspended solid (TSS; 0 mg/L, 3,000 mg/L) treatment groups sampled during the first 36 h postfertilization at Tishomingo National Fish Hatchery in Tishomingo, Oklahoma, on 26–27 July 2012. Developmental stages include 512-cell stage (512-S), late blastula (LB), gastrula (G), neurula (N), blastopore closure (BC), somite appearance (SA), optic primordium (OP), tail bud (TB), and otolith appearance (OA).

Found at DOI: http://dx.doi.org/10.3996/112015-JFWM-111.S1 (4 KB TXT).

Reference S1. Chapman DC, editor. 2007. Early development of four cyprinids native to the Yangtze River, China. U.S. Geological Survey Data Series 239.

Found at DOI: http://dx.doi.org/10.3996/112015-JFWM-111.S2; also available at http://pubs.usgs.gov/ds/2006/239/pdf/ds2006-239.pdf (2,863 KB PDF).

Reference S2. Cowley DE, Alleman J, Mcshane R, Shirey P, Sallenave R. 2005. Effects of sediment and salinity on physical properties of the egg of the Rio Grande Silvery Minnow (Hybognathus amarus). New Mexico Water Resources Research Institute, Technical Completion Report 334.

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Reference S3. Fields CB, Mortsch LD, Brklacich M, Forbes DL, Kovacs P, Patz JA, Running SW, Scott MJ. 2007. North American. Climate change 2007: impacts, adaptation and vulnerability. Pages 617–652 in Parry ML, Canziani OF, Palutikof JP, Van der Linden PJ, Hanson CE, editors. Contribution of Working Group II to the fourth assessment report of the Intergovernmental Panel on Climate Change. Cambridge, England, UK: Cambridge University Press.

Found at DOI: http://dx.doi.org/10.3996/112015-JFWM-111.S4; also available at https://www.ipcc.ch/pdf/assessment-report/ar4/wg2/ar4-wg2-chapter14.pdf (872 KB PDF).

Reference S4. Jones P, Martin F, Hardy J Jr. 1978. Development of fishes of the mid-Atlantic bight, volume 1. U. S. Fish and Wildlife Service Biological Services Program FWS. OBS-78/12.

Found at DOI: http://dx.doi.org/10.3996/112015-JFWM-111.S5; also available at http://docs.lib.noaa.gov/rescue/NOAA_E_DOCS/Coastal_Services_Center_docs/Development%20of%20Fishes%20of%20the%20Mid-Atlantic%20Bight%20-%20An%20Atlat%20of%20Eg/ Volume%20I%20-%20Acipenseridae%20through% 20Ictaluridae.pdf (20,072 KB PDF).

Reference S5. Mueller JS. 2013. Effects of temperature, salinity, and suspended solids on the early life history stages of Arkansas River Shiner. Master's thesis. Texas Tech University, Lubbock, Texas.

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We thank D. Fenner, B. Bristow, W.H. Brandenbug, A.L. Barkalow, S.A. Zipper, and S.P. Platania for providing the Arkansas River Shiners used in this study and K. Graves and the staff of the Tishomingo National Fish Hatchery for providing facilities and technical expertise and advice for spawning Arkansas River Shiner. G. Wilde, the Associate Editor, and three anonymous reviewers provided comments and suggestions that greatly improved an earlier version of this manuscript. This study was performed under the auspices of Texas Tech University (AUP #11061-08) and U.S. Fish and Wildlife Service Federal Fish and Wildlife Permit #TE48766A-0.

Funding was provided by the U.S. Fish and Wildlife Service Great Plains Landscape Conservation Cooperative (U.S. Fish and Wildlife Service agreement F11AP00112). Cooperating agencies for the Texas Cooperative Fish and Wildlife Research Unit are the U.S. Geological Survey, Texas Tech University, Texas Parks and Wildlife, and the Wildlife Management Institute. Cooperating agencies for the Oklahoma Cooperative Fish and Wildlife Research Unit are the U.S. Geological Survey, Oklahoma State University, and the Oklahoma Department of Wildlife Conservation and the Wildlife Management Institute. Cooperating agencies for the Hawaii Cooperative Fishery Research Unit are the U.S. Geological Survey, the University of Hawaii system, the Hawaii Department of Land and Natural Resources, and the U.S. Fish and Wildlife Service.

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