Michal B&#237;l&#253; and Ond&#345;ej P. Simon were supported by grants from the Czech University of Life Science [Internal Grant Agency of Faculty of Environmental Sciences, CULS Prague (42110 1312 3175 (20164236))]. Support for Karel Douda came from the Czech Science Foundation (13-05872S). Data on the bioindication and present occurrence of pearl mussels were collected during the implementation of the Czech Action Plan for Freshwater Pearl Mussels managed by the Nature Conservation Agency of the Czech Republic, which is funded by the government of the Czech Republic and is available at

1. Mesh Cage
Note: See Figure 1.
<ol>
	<li>Prepare material
	<ol>
		<li>Prepare the material for the in-laboratory part of the experiment: ~1 - 2 L of river water per mesh cage, mesh cages (1 main plastic body, 2 plastic covers, 2 sheets of special technical sieves with 340 &#181;m pores, 4 bolts and 4 nuts per cage), pliers, a spanner, Pasteur pipettes, a strainer, a digital camera, a trinocular dissecting zoom stereo microscope, a calibration grid (microscope equipment), 5 Petri dishes of 50 mm diameter, beakers, 2 plastic dishes (~25 cm x 15 cm x 3 - 5 cm), and a plastic box.</li>
		<li>To perform the hyporheal installation, prepare a rubber hose and a 100-&#181;m-pore mesh, and a squirt bottle. For the construction of the device, see Supplementary File 1: S.1. Mesh cages construction.</li>
	</ol>
	</li>
	<li>Assemble the bottom and central part of the mesh cages. Assemble the part of the cage that holds the individuals. Insert one plastic cover first, then one sheet of the plastic sieve, and finally the main body on top. Use four bolts to secure it.</li>
	<li>Prepare biological material 
	<ol>
		<li>Put the mesh cage into the plastic dish containing river water. Ensure that the chambers are half full. Take the FWPM juveniles (see Supplementary File 1: S.6. Biological material) out of the thermally-insulated box and put them in the Petri dish. 
		Note: Ensure that sudden temperature changes do not exceed ~2 &#176;C.</li>
		<li>Using a squirt bottle and strainer, sift through the juveniles to clear the detritus.</li>
	</ol>
	</li>
	<li>Set up the microscope and camera. Perform a calibration of the instruments (see Supplementary File1: S. 5. Microscope and phototechnics). Place a Petri dish containing a little water under the microscope.</li>
	<li>Put the juveniles into cages (experimental laboratory work)
	<ol>
		<li>Use a Pasteur pipette to remove one individual from a Petri dish and carefully place it in the Petri dish under the microscope.</li>
		<li>Check the individual&#8217;s fitness by looking into the eyepiece (~40X magnification). 
		Note: &#8220;Good&#8221; fitness signifies that the individual moves, rotates from side to side, pushes the foot out of the shell, etc. Remove dead or low fitness individuals with a Pasteur pipette and place them in a separate Petri dish (FWPM juveniles with an opened shell, no movement, the foot is not pulled out, a fragmented shell, juveniles who float uncontrollably in the water, a visible decomposition of the shell, partial decalcification).</li>
		<li>Take two photographs of an FWPM individual showing good fitness using a constant magnification of ~80X. See Supplementary File 1: S.5. Microscope and phototechnics. Save the photos. 
		Note: For a good measurement of its length, the juvenile must be laid lengthwise (lateral view). The main goal is to take a high-quality picture of the maximum shell length good enough to enable a picture analysis afterward.</li>
		<li>Insert the juvenile into the appropriate chamber in the cage as soon as the pictures are taken. Record the numbers of the pictures and the chamber.</li>
		<li>Repeat this step with each individual for all the used chambers in the mesh cage. 
		Note: see Supplementary File 1: S.1. Mesh cages construction.</li>
		<li>Once all the used chambers have pearl mussels, put the plastic sieve on the cage, then gently put the plastic cover on and secure all parts together with the nuts.</li>
		<li>In the case of an installation into a hyporheic zone, pass one of the hose ends through one of the chambers and fix it in this position, then take the anti-clogging mesh and bind it on the bottom end (see Supplementary File 1: S.1. Mesh cages construction).</li>
	</ol>
	</li>
	<li>Store juveniles 
	<ol>
		<li>Put the cage into the plastic box with the river water, so that the juveniles are fully immersed, and keep it in the thermobox. Before the installation, let the juveniles adapt to the in situ river water temperature at the place of installation (gradual cooling, max. 5 &#176;C in 24 h).</li>
	</ol>
	</li>
	<li>Install mesh cages
	<ol>
		<li>Prepare the field material including the mesh cages with the juveniles, steel spikes, bolts and metal nuts, a spanner, field temperature dataloggers (see Table of Materials and Supplementary File 1: S.4.2. Water measurement), a string, a camera, the field protocol, a hammer, and a spade.</li>
		<li>Transport the FWPM juveniles to the site in a field thermobox (insulated box), keeping a stable water temperature with variations &#60; ~2 &#176;C. Put the thermobox with the mesh cages into the river on the site to let the juveniles adapt to the local environmental conditions (pH, conductivity, etc.).</li>
		<li>Install the mesh cage.
		<ol>
			<li>Remove the mesh cage from the field thermobox. Provide it with two steel spikes and fasten the field datalogger. Anchor the cage into a habitat with conditions typical for FWPMs in the study area (e.g., at the edge of the main stream flow, not in direct water flow, not in standing water, not in direct sunlight).
			<ol>
				<li>For open water, using a pair of the steel spikes, fix the cage to the river bottom; lay it on its side and level with the river bottom, downstream at an angle of 45&#176; to the river flow, towards the center of the river. The lower horizontal edge should be about 10 - 15 cm above the river bottom surface. Maintain a minimum distance of 2 m between each cage at one locality (see Supplementary File 1: S.4. Cages maintenance).</li>
				<li>For the hyporheic zone, dig the cages into the river bottom in a perpendicular landscape position, perpendicular to the stream of water, so that the upper horizontal edge of the cage is parallel to the river bottom surface and the chambers are located at the hyporheic depth which should be tested. Take out the upper end of the rubber hose above the bottom surface for the possibility of water sampling during the experiment (see Supplementary File 1: S.4.2. Water measurement). 
				Note: It is recommended to perform regular checks and maintenance on the cages (see Supplementary File 1: S. 4. Cages maintenance).</li>
			</ol>
			</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Uninstall the cages and transport the juveniles after the exposure. For this, pull the cages out of the water, clear them of fine sediment as well as of drifted material and put them into the field thermobox filled with river water. Transport the cages immediately to the laboratory and start the mortality and growth rate evaluation. 
	Note: See Supplementary File 1: S.3. Exposure duration. In the case of a temperature difference of more than 5 &#176;C between the cages and the laboratory environment, it is first necessary to let the temperature equalize.</li>
	<li>Evaluate the experiment by checking the life/fitness of each juvenile (see steps 1.5.2 and 1.5.3) and take 2 images of each live juvenile in a Petri dish using a constant magnification of ~80X. Record the fitness and the numbers of the pictures and chambers.</li>
	<li>Complete the experiment (common to all methods)
	<ol>
		<li>Perform the measurements in image analysis software. Use image analysis software for the body size determination of every evaluated juvenile on both the input images (step 1.5.3) and on the output images (step 1.9). Use the maximum total shell length recorded in both photographs as body size values in both input and output.</li>
		<li>Insert the measured values into the table processor and calculate the growth increment (%) for each surviving juvenile.</li>
		<li>Estimate the survival rate (%) per mesh cage using the ratio of the number of surviving individuals to all experimental individuals in the mesh cage. 
		Note: After the experiment, return the survivors to the breeding program 
		(see Supplementary File 1: S.6. Biological material).</li>
	</ol>
	</li>
</ol>
2. Sandy Cage
Note: See Figure 2.
<ol>
	<li>Prepare material 
	<ol>
		<li>Prepare the material for the in-laboratory part of the experiment: 2 Petri dishes (diameter ~8.5 cm), Pasteur pipettes, a strainer, 25 L of river water, a plastic box, sieves (mesh size 1 and 2 mm), a big plastic box (25 L), a sandy cage (see Supplementary File 1: S.2. Sandy cages construction), a digital camera, a trinocular dissecting zoom stereo microscope, a calibration grid (microscope equipment), sorted river sand from the study area (see step 2.1.3), and the protocol. See Table of Materials and Supplementary File 1: S. 2. Sandy cages construction.</li>
		<li>Prepare the material for the isolation process: round containers (1 for each cage plus 1 extra), 2 Petri dishes (diameter ~14 cm), a Pasteur pipette, magnifying glasses, and 1 L of river water.</li>
		<li>Sift the river sand through a 2-mm sieve and then through a 1-mm sieve to get a grain size of 1 - 2 mm. Dry the sand and save it in a dry form until required.</li>
	</ol>
	</li>
	<li>Take the juveniles (see Supplementary File 1: S.6. Biological material) out of the thermobox and put them in the Petri dish. Using a squirt bottle and strainer, sift through the juveniles to clear the detritus.</li>
	<li>Set up the microscope and camera (see Supplementary File 1: S.5. Microscope and phototechnics).</li>
	<li>Put juveniles into cages (experimental laboratory work)
	<ol>
		<li>Place the sandy cage in the plastic box. Scatter the sorted sand (see step 2.1.3) up to one-third of the height of the sandy cage. Pour water into the box. Ensure that the sand surface is about 10 mm below the water level. Insert the sandy cage into the 25 L box of river water and expose it to the same temperature as the juvenile FWPMs (see Supplementary File 1: S.6.2. Storage of the biological material) for 12 h. Avoid any exposure of the sand to sunlight.</li>
		<li>Take the Petri dish with the prepared FWPM juveniles.</li>
		<li>Check the individuals&#8217; fitness by looking into the eyepiece (see step 1.5.2).</li>
		<li>Perform the photographic documentation as follows. Take one picture of all individuals discovered (see step 1.5.3) and choose 10 of the largest individuals. Alternatively, take pictures of all juveniles together with low magnification (~40X) for a bulk evaluation and choose the 10 largest individuals. Save all the photos and record their numbers.</li>
		<li>Using a squirt bottle, move the FWPM juveniles into the prepared sandy cage.</li>
	</ol>
	</li>
	<li>Store juveniles 
	<ol>
		<li>Put the cage into the big plastic box with river water so that the cage is fully immersed and keep it in the thermobox. Let the juveniles adapt to the in situ river water temperature (gradual cooling, max. 5 &#176;C for 24 h) before the installation.</li>
	</ol>
	</li>
	<li>Install sandy cages
	<ol>
		<li>Prepare the material for the field installation: sandy cages, a ~25-L field thermobox, a flat stone (minimal weight 1 kg), a net (mesh size 10 x 10 mm), a squirt bottle, field temperature dataloggers (see Table of Materials and Supplementary File 1: S.4.2. Water measurement), a spade, and the field protocol.</li>
		<li>Transport the cages with the juveniles to the site in the field thermobox, keeping a stable water temperature (~2 &#176;C change). Put the field thermobox with the sandy cages into the river at the field site to let the FWPM juveniles adapt to the local environmental conditions (pH, conductivity, etc.).</li>
		<li>Install the sandy cages into habitats with conditions typical for FWPMs (e.g., at the edge of the main stream flow in a meander, not in direct water flow, not in standing water, not in direct sunlight).
		<ol>
			<li>For open water, fasten the sandy cages to a flat stone using a net and place it on the river bottom. Ensure that the larger side of the cage forms an angle of 45&#176; with the flow.</li>
			<li>For Hyporheal, dig the cages into the river bottom perpendicular to the flow of water so that the cage lid is level with the river bottom surface. 
			Note: It is recommended to perform regular checks and maintenance on the cages (see Supplementary File 1: S. 4. 1. Site checks).</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Uninstall cages and transport juveniles after exposure 
	Note: see Supplementary File 1: S.3. Exposure duration.
	<ol>
		<li>Pull the cages out of the water, clear them of drifted material and put them into the field thermobox filled with river water.</li>
		<li>Transport the cages to the laboratory and start the mortality and growth rate evaluation. 
		Note: In the case of a temperature difference of more than 5 &#176;C between the cages and the laboratory environment, it is necessary to let the temperatures equalize.</li>
	</ol>
	</li>
	<li>Separate FWPM juveniles from sand
	<ol>
		<li>Prepare a round container with a water depth of 50 mm (for each cage separately) and one extra round container. Transfer the sand from the cage into the round container. Use a swirling motion to wash out the lighter particles into an extra container.</li>
		<li>Sample the content from this container gradually and search for juveniles step-by-step using a Pasteur pipette and a magnifying glass. Put the juveniles in the Petri dish using the Pasteur pipette. Repeat this step until the last juvenile has been found and then another 10x after the first negative finding. After each wash step, add clean river water to the original container with sand. 
		Note: Especially after the first washing out, properly examine the content and clean it of ballast such as fine sediment and other alluvia.</li>
	</ol>
	</li>
	<li>Evaluate the experiment
	<ol>
		<li>Check the fitness of each juvenile (see steps 2.4.3 and 1.5.2) and count the number of survivors.</li>
		<li>Take a picture (see step 2.4.4.) of each individual separately, although this means there is no clear identity of each individual. Alternatively, take bulk photos and choose a subset of the 10 best-grown individuals from the final results. 
		Note: Both possibilities have a similar reporting value. Possibility 1 has a limitation of a higher workload but also the benefit of the highest photo magnification and thus also greater accuracy.</li>
	</ol>
	</li>
	<li>Complete the experiment
	<ol>
		<li>Perform measurements in image analysis software. Complete the experiment as done in the mesh cages (see step 1.10) with the following exception: do not evaluate the growth rate (%) of each juvenile but evaluate the group as a whole in the sandy cage experiment. 
		Note: After the experiment, the survivors should be returned to the breeding program 
		(see Supplementary File S.6.1. Selection of a biological material).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Exposure time:
Even one-month exposed mesh cages show a visible growth reflecting differences between localities (Figure 3), so they are very usable for the quick and easy detection of a locality characterization. Nevertheless, the relevance of the results depends on the short-term state of the conditions, which can oscillate. In particular, short rainfall events can play a role. In contrast, unpredictable episodic pollution may not always be recorded. In locality V ...

The exposure of experimental organisms in situ with the subsequent evaluation of their condition is one possible way to get information about the environmental quality and (especially) the site suitability for a species. Within animals, such a bioindication is applicable primarily for small invertebrates which are able to live in a limited bounded space. Young stages of bivalves (Bivalvia) are one such suitable organism group1.
Bivalves of the family Unionidae are a very important component of aquatic ecosystems2. However, these species are often critically endangered, especially in streams and rivers. Some of them are characterized as &#39;umbrella species&#39; whose conservation is closely related to the conservation of the whole stream biotope and which require a comprehensive approach3. These animals have a life cycle associated with many environment components, from water chemistry4,5 to changes in the populations of fish which serve as mussel larvae hosts6. Because mussel juveniles often represent a critical phase of the mussel life cycle, the site suitability for their development at this stage is crucial for a successful species population development in a locality.
The freshwater pearl mussel (FWPM, Margaritifera margaritifera; Unionida, Bivalvia) is a critically endangered bivalve occurring in oligotrophic European streams. Their numbers have fallen drastically during the 20th century across the occurrence area. It seems that the current decline in species reproduction in the majority of the central European populations is primarily caused by very low to zero survival of juveniles during the first few years of their life. It is assumed that juvenile FWPMs live for many years in the shallow hyporheic zone7, of which the conditions and their variability are still not well described. Moreover, until their second year of life, the juveniles only have a dimension of up to about 1 mm, so they are very difficult to find in large volumes of sediment under natural conditions8. Therefore, experiments with captive juveniles are necessary for the study of their ecology.
Within the Czech Action Plan for Freshwater Pearl Mussel9, there are thousands of juveniles rising every year from a semi-natural breeding program. Nevertheless, there is a question of which localities and habitats are suitable for successful population support by these juveniles or for eventual species reintroduction. In situ bioindications present a way of finding the answer.
Despite the fact that inconsistent survival rates of juvenile mussels in exposure cages were observed in some earlier works that questioned the suitability of juvenile mussels as bioindicators10, several recent studies have confirmed the applicability of juvenile exposure methods for water quality testing11,12,13. Additionally, it has been demonstrated that several factors need to be considered when interpreting the results of these particular studies, such as the stock origin14 and the persisting effects of larval conditions15.
The question arises of how to install experimental juveniles in tested localities and how to most effectively evaluate their condition. The first rigorous application of in situ exposure methods with juvenile FWPMs was published by Buddensiek16. Juvenile FWPM individuals were kept in sheet cages, exposed in the free-flowing water of streams, and their survival and growth were quantified after several weeks of exposure. The approach was originally developed as a semi-artificial breeding method, but the author also highlighted its applicability for the assessment of habitat requirements and water quality. Although the FWPM juvenile survival is naturally very low on a scale of months/years and only a very small number of animals will survive, the survival rate can be a good marker of the environmental effect on a scale of several weeks16. Over years of research, exposure methods were developed further to hold experimental juvenile mussel in-stream habitats and to evaluate their growth and survival rates; these include sandy boxes17, mussel silos based on an upwelling principle18, and various other exposure cages (summarized by Gum and colleagues)11. Because juveniles occur naturally in shallow hyporheic zone7, the application of experimental devices within the stream bottom is very desirable.
In our article, we describe the use of two exposure devices for FWPMs: i) modified Buddensiek sheet cages (&#34;mesh cages&#34;) also enabling bioindication testing in hyporheal conditions; and ii) Hru&#353;ka sandy boxes (&#34;sandy cages&#34;). The protocol describes the application of both methods in open water and hyporheic conditions (i.e., four variants of exposure are described). The methods were gradually modified and expanded over more than 15 years of application within the Czech Action Plan for Freshwater Pearl Mussel9 and verified by a set of experiments.

<table><tbody><tr><td>&lt;strong&gt;biological material maintenance and care&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Freshwater pearl mussel juveniles</td><td>any</td><td>NA</td><td>from a FWPM breeding programme</td></tr><tr><td>plastic boxes</td><td>any</td><td>NA</td><td></td></tr><tr><td>thermobox</td><td>MERCI</td><td>212,070,600,030</td><td>There are many possibilities. This is one example only.</td></tr><tr><td>field thermobox (ca25 l)</td><td>any</td><td>NA</td><td>cold box (insulated box) commonly used for food transport</td></tr><tr><td>river water</td><td>any</td><td>NA</td><td></td></tr><tr><td>Petri dishes</td><td>any</td><td>NA</td><td></td></tr><tr><td>plastic Pasteur pipettes with balloon bulb (droppers)</td><td>any</td><td>NA</td><td>hole diameter 1 mm</td></tr><tr><td>hydrogen peroxide</td><td>any</td><td>NA</td><td></td></tr><tr><td>plastic container (ca 50 l) for river water</td><td>any</td><td>NA</td><td></td></tr><tr><td>plastic tea strainer</td><td>any</td><td>NA</td><td>commonly used in kitchen</td></tr><tr><td>&lt;strong&gt;mesh cages construction&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>main plastic bodies</td><td>any</td><td>NA</td><td></td></tr><tr><td>plactic covers</td><td>any</td><td>NA</td><td></td></tr><tr><td>special technical sieves 340 &amp;micro;m</td><td>Silk &amp;amp;Progress</td><td>UHELON 20 T</td><td></td></tr><tr><td>special technical sieves 100 &amp;micro;m</td><td>Silk &amp;amp;Progress</td><td>UHELON 67 M</td><td></td></tr><tr><td>rubber hose (diameter 5.5 mm)</td><td>any</td><td>NA</td><td></td></tr><tr><td>steel bolts</td><td>any</td><td>NA</td><td></td></tr><tr><td>steel nuts</td><td>any</td><td>NA</td><td></td></tr><tr><td>spanner</td><td>any</td><td>NA</td><td></td></tr><tr><td>steel spikes</td><td>any</td><td>NA</td><td></td></tr><tr><td>pliers</td><td>any</td><td>NA</td><td></td></tr><tr><td>beakers</td><td>any</td><td>NA</td><td></td></tr><tr><td>plastic dishes (ca. 25x15x3-5cm)</td><td>any</td><td>NA</td><td></td></tr><tr><td>squirt bottle</td><td>any</td><td>NA</td><td></td></tr><tr><td>field protocols</td><td>any</td><td>NA</td><td></td></tr><tr><td>stationery</td><td>any</td><td>NA</td><td></td></tr><tr><td>plastic container</td><td>any</td><td>NA</td><td></td></tr><tr><td>string</td><td>any</td><td>NA</td><td></td></tr><tr><td>hammer</td><td>any</td><td>NA</td><td></td></tr><tr><td>&lt;strong&gt;sandy cages construction and use&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>sieve 1 mm</td><td>any</td><td>NA</td><td></td></tr><tr><td>sieve 2 mm</td><td>any</td><td>NA</td><td></td></tr><tr><td>special technical sieves 340 &amp;micro;m</td><td>Silk &amp;amp;Progress</td><td>UHELON 20 T</td><td></td></tr><tr><td>plastic boxes with tight-fitting lid</td><td>any</td><td>NA</td><td></td></tr><tr><td>hot melt adhesive</td><td>any</td><td>NA</td><td></td></tr><tr><td>plastic box (ca 250 x 150 x 100 cm)</td><td></td><td></td><td></td></tr><tr><td>big plastic box (ca 25 l)</td><td>any</td><td>NA</td><td></td></tr><tr><td>flat stone</td><td>any</td><td>NA</td><td></td></tr><tr><td>net</td><td>any</td><td>NA</td><td></td></tr><tr><td>river sand</td><td>any</td><td>NA</td><td></td></tr><tr><td>round containers</td><td>any</td><td>NA</td><td></td></tr><tr><td>magnifying glasses</td><td>Carson</td><td>Carson CP 60</td><td>There ar many possibilities. This is one example only</td></tr><tr><td>&lt;strong&gt;cages installation and maintenance&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>field temperature dataloggers</td><td>ONSET</td><td>UA-001-64</td><td>http://www.onsetcomp.com/products/data-loggers/ua-001-64</td></tr><tr><td>spade</td><td>any</td><td>NA</td><td></td></tr><tr><td>toothbrush</td><td>any</td><td>NA</td><td></td></tr><tr><td>&lt;strong&gt;experiment evaluation&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>trinocular dissecting zoom stereo microscope</td><td>Bresser optic</td><td>ICD 10x-160x</td><td>There are many possibilities. This is one example only.</td></tr><tr><td>digital camera/ electronic eyepiece</td><td>Bresser optic</td><td>MikroCamLab 5M</td><td>There are many possibilities. This is one example only.</td></tr><tr><td>Calibration gird</td><td>Am Scope</td><td>SKU: MR100</td><td>There are many possibilities. This is one example only.</td></tr><tr><td>external power source with two movable light guides</td><td>Arsenal</td><td>K1309010150021</td><td>There are many possibilities. This is one example only.</td></tr><tr><td>Image software</td><td>ImageJ software</td><td></td><td>There are many possibilities. This is one example only.</td></tr><tr><td>table processor</td><td>MS excel</td><td></td><td>There are many possibilities. This is one example only.</td></tr></tbody></table>

bioindication testing of stream environment suitability for young freshwater pearl mussels using in situ exposure methods

Knowledge of habitat suitability for freshwater mussels is an important step in the conservation of this endangered species group. We describe a protocol for performing in situ juvenile exposure tests within oligotrophic river catchments over one-month and three-month periods. Two methods (in both modifications) are presented to evaluate the juvenile growth and survival rate. The methods and modifications differ in value for the locality bioindication and each has its benefits as well as limitations. The sandy cage method works with a large set of individuals, but only some of the individuals are measured and the results are evaluated in bulk. In the mesh cage method, the individuals are kept and measured separately, but a low individual number is evaluated. The open water exposure modification is relatively easy to apply; it shows the juvenile growth potential of sites and can also be effective for water toxicity testing. The within-bed exposure modification needs a high workload but is closer to the conditions of a natural juvenile environment and it is better for reporting the real suitability of localities. On the other hand, more replications are needed in this modification due to its high-hyporheic environment variability.

The four bioindication methods (open water sandy cages, within-bed sandy cages, open water mesh cages, and within-bed mesh cages) were applied to investigate the environment condition suitability for FWPMs in the upper Vltava River Basin (Bohemian Forest, Czech Republic). This river represents one FWPM residual locality within central Europe19. Here, we present a specially selected set of results illustrating the most important aspects of the four methods. Further ...

Watch this Scientific Journal Video about Bioindication Testing of Stream Environment Suitability for Young Freshwater Pearl Mussels Using In Situ Exposure Methods at JoVE.com

Bioindication Testing of Stream Environment Suitability for Young Freshwater Pearl Mussels Using In Situ Exposure Methods

In situ bioindications enable determination of the suitability of an environment for endangered mussel species. We describe two methods based on the juvenile exposure of freshwater pearl mussels in cages to oligotrophic river habitats. Both methods are implemented in variants for open water and hyporheic water environments.

Bioindication Testing of Stream Environment Suitability for Young Freshwater Pearl Mussels Using In Situ Exposure Methods

Knowledge of habitat suitability for freshwater mussels is an important step in the conservation of this endangered species group. We describe a protocol ...

bioindication-testing-stream-environment-suitability-for-young

Research

JoVE Journal

Environment

7.3K Views.  Czech University of Life Sciences Prague. In situ bioindications inschakelen bepaling van de geschiktheid van een omgeving voor bedreigde Mossel soorten. Beschrijven we twee methoden op basis van de jonge Gasbedwelming met behulp van zoetwater parel mosselen in kooien oligotrofe rivierhabitats. Beide methoden zijn geïmplementeerd in de varianten voor open water en hyporheic water omgevingen.

Video: Bioindication Testing of Stream Environment Suitability for Young Freshwater Pearl Mussels Using In Situ Exposure Methods

Use of Chironomidae (Diptera) Surface-Floating Pupal Exuviae as a Rapid Bioassessment Protocol for Water Bodies

Rapid bioassessment protocols using benthic macroinvertebrate assemblages have been successfully used to assess human impacts on water quality. Unfortunately, traditional benthic larval sampling methods, such as the dip-net, can be time-consuming and expensive. An alternative protocol involves collection of Chironomidae surface-floating pupal exuviae (SFPE). Chironomidae is a species-rich family of flies (Diptera) whose immature stages typically occur in aquatic habitats. Adult chironomids emerge from the water, leaving their pupal skins, or exuviae, floating on the water&#8217;s surface. Exuviae often accumulate along banks or behind obstructions by action of the wind or water current, where they can be collected to assess chironomid diversity and richness. Chironomids can be used as important biological indicators, since some species are more tolerant to pollution than others. Therefore, the relative abundance and species composition of collected SFPE reflect changes in water quality. Here, methods associated with field collection, laboratory processing, slide mounting, and identification of chironomid SFPE are described in detail. Advantages of the SFPE method include minimal disturbance at a sampling area, efficient and economical sample collection and laboratory processing, ease of identification, applicability in nearly all aquatic environments, and a potentially more sensitive measure of ecosystem stress. Limitations include the inability to determine larval microhabitat use and inability to identify pupal exuviae to species if they have not been associated with adult males.

Use of Chironomidae Diptera Surface-Floating Pupal Exuviae as a Rapid Bioassessment Protocol for Water Bodies

Rapid bioassessment protocols using benthic macroinvertebrates are often used to monitor and assess water quality. An efficient protocol involves collections of Chironomidae surface-floating pupal exuviae (SFPE). Here, techniques for field collection, laboratory processing, slide mounting, and identification of Chironomidae SFPE are described.

Rapid bioassessment protocols using benthic macroinvertebrate assemblages have been successfully used to assess human impacts on water quality. ...

Use of a Battery of Chemical and Ecotoxicological Methods for the Assessment of the Efficacy of Wastewater Treatment Processes to Remove Estrogenic Potency

Endocrine Disrupting Compounds pose a substantial risk to the aquatic environment. Ethinylestradiol (EE2) and estrone (E1) have recently been included in a watch list of environmental pollutants under the European Water Framework Directive. Municipal wastewater treatment plants are major contributors to the estrogenic potency of surface waters. Much of the estrogenic potency of wastewater treatment plant (WWTP) effluents can be attributed to the discharge of steroid estrogens including estradiol (E2), EE2 and E1 due to incomplete removal of these substances at the treatment plant. An evaluation of the efficacy of wastewater treatment processes requires the quantitative determination of individual substances most often undertaken using chemical analysis methods. Most frequently used methods include Gas Chromatography-Mass Spectrometry (GCMS/MS) or Liquid Chromatography-Mass Spectrometry (LCMS/MS) using multiple reaction monitoring (MRM). Although very useful for regulatory purposes, targeted chemical analysis can only provide data on the compounds (and specific metabolites) monitored. Ecotoxicology methods additionally ensure that any by-products produced or unknown estrogenic compounds present are also assessed via measurement of their biological activity. A number of in vitro bioassays including the Yeast Estrogen Screen (YES) are available to measure the estrogenic activity of wastewater samples. Chemical analysis in conjunction with in vivo and in vitro bioassays provides a useful toolbox for assessment of the efficacy and suitability of wastewater treatment processes with respect to estrogenic endocrine disrupting compounds. This paper utilizes a battery of chemical and ecotoxicology tests to assess conventional, advanced and emerging wastewater treatment processes in laboratory and field studies.

Endocrine Disrupting Compounds (EDC) pose a substantial risk to the aquatic environment. Municipal wastewater treatment plants are major contributors to the estrogenic potency of surface waters. The methodology provided in this paper allows for an assessment of the efficacy and suitability of wastewater treatment processes with respect to EDC removal.

Endocrine Disrupting Compounds pose a substantial risk to the aquatic environment. Ethinylestradiol (EE2) and estrone (E1) have recently been included in ...

Salinity-dependent Toxicity Assay of Silver Nanocolloids Using Medaka Eggs

Salinity is an important characteristic of the aquatic environment. For aquatic organisms it defines the habitats of freshwater, brackish water, and seawater. Tests of the toxicity of chemicals and assessments of their ecological risks to aquatic organisms are frequently performed in freshwater, but the toxicity of chemicals to aquatic organisms depends on pH, temperature, and salinity. There is no method, however, for testing the salinity dependence of toxicity to aquatic organisms. Here, we used medaka (Oryzias latipes) because they can adapt to freshwater, brackish water, and seawater. Different concentrations of embryo-rearing medium (ERM) (1x, 5x, 10x, 15x, 20x, and 30x) were employed to test the toxicity of silver nanocolloidal particles (SNCs) to medaka eggs (1x ERM and 30x ERM have osmotic pressures equivalent to freshwater and seawater, respectively). In six-well plastic plates, 15 medaka eggs in triplicate were exposed to SNCs at 10 mg/L&#8722;1 in different concentrations of ERM at pH 7 and 25 &#176;C in the dark.
We used a dissecting microscope and a micrometer to measure heart rate per 15 sec and eye diameter on day 6 and full body length of the larvae on hatching day (section 4). The embryos were observed until hatching or day 14; we then counted the hatching rate every day for 14 days (section 4). To see silver accumulation in embryos, we used inductively coupled plasma mass spectrometry to measure the silver concentration of test solutions (section 5) and dechorionated embryos (section 6).The toxicity of the SNCs to medaka embryos obviously increased with increasing salinity. This new method allows us to test the toxicity of chemicals in different salinities.

Embryonic stages are the most susceptible to xenobiotics. Although chemical toxicity depends on salinity, no method exists to test the salinity dependence of toxicity to aquatic organisms. Here, we describe a new and high-throughput method for determining the salinity dependence of toxicity to aquatic embryos.

Salinity is an important characteristic of the aquatic environment. For aquatic organisms it defines the habitats of freshwater, brackish water, and ...

Developmental Biology

Ecotoxicological Method with Marine Bacteria Vibrio anguillarum to Evaluate the Acute Toxicity of Environmental Contaminants

Bacteria are an important component of the ecosystem, and microbial community alterations can have a significant effect on biogeochemical cycling and food webs. Toxicity testing based on microorganisms are widely used because they are relatively quick, reproducible, cheap, and are not associated with ethical issues. Here, we describe an ecotoxicological method to evaluate the biological response of the marine bacterium Vibrio anguillarum. This method assesses the acute toxicity of chemical compounds, including new contaminants such as nanoparticles, as well as environmental samples. The endpoint is the reduction of bacterial culturability (i.e., the capability to replicate and form colonies) due to exposure to a toxicant. This reduction can be generally referred to as mortality. The test allows for the determination of the LC50, the concentration that causes a 50% decrease of bacteria actively replicating and forming colonies, after a 6 h exposure. The culturable bacteria are counted in terms of colony forming units (CFU), and the &#34;mortality&#34; is evaluated and compared to the control. In this work, the toxicity of copper sulphate (CuSO4) was evaluated. A clear dose-response relationship was observed, with a mean LC50 of 1.13 mg/L, after three independent tests. This protocol, compared to existing methods with microorganisms, is applicable in a wider range of salinity and has no limitations for colored/turbid samples. It uses saline solution as the exposure medium, avoiding any possible interferences of growth medium with the investigated contaminants. The LC50 calculation facilitates comparisons with other bioassays commonly applied to ecotoxicological assessments of the marine environment.

Ecotoxicological Method with Marine Bacteria Vibrio anguillarum to Evaluate the Acute Toxicity of Environmental Contaminants

This work describes a new protocol to assess the ecotoxicity of pollutants, including emerging contaminants such as nanomaterials, using the marine bacterium Vibrio anguillarum. This method allows for the determination of the LC50, or mortality, the concentration that causes a 50% decrease in bacteria culturability, after a 6 h exposure.

Bacteria are an important component of the ecosystem, and microbial community alterations can have a significant effect on biogeochemical cycling and food ...

Protocol for Acute and Chronic Ecotoxicity Testing of the Turquoise Killifish Nothobranchius furzeri

The killifish Nothobranchius furzeri is an emerging model organism in the field of ecotoxicology and its applicability in acute and chronic ecotoxicity testing has been demonstrated. Overall, the sensitivity of the species to toxic compounds is in the range with, or higher than, that of other model species.
This work describes protocols for acute, chronic, and multigenerational bioassays of single and combined stressor effects on N. furzeri. Due to its short maturation time and life-cycle, this vertebrate model allows the study of endpoints such as maturation time and fecundity within four months. Transgenerational full life-cycle exposure trials can be performed in as little as 8 months. Since this species produces eggs that are drought-resistant and remain viable for years, the on-site culture of the species is not needed but individuals can be recruited when required. The protocols are designed to measure life-history traits (mortality, growth, fecundity, weight) and critical thermal maximum.

Protocol for Acute and Chronic Ecotoxicity Testing of the Turquoise Killifish Nothobranchius furzeri

In this work, we describe an acute, chronic and multigenerational bioassay to study the effects of single and combined stressors on the Turquoise killifish Nothobranchius furzeri. This protocol is designed to study life-history traits (mortality, growth, fecundity, weight) and critical thermal maximum.

The killifish Nothobranchius furzeri is an emerging model organism in the field of ecotoxicology and its applicability in acute and chronic ecotoxicity ...

Design and Use of an Apparatus for Quantifying Bivalve Suspension Feeding at Sea

As shellfish aquaculture moves from coastal embayments and estuaries to offshore locations, the need to quantify ecosystem interactions of farmed bivalves (i.e., mussels, oysters, and clams) presents new challenges. Quantitative data on the feeding behavior of suspension-feeding mollusks is necessary to determine important ecosystem interactions of offshore shellfish farms, including their carrying capacity, the competition with the zooplankton community, the availability of trophic resources at different depths, and the deposition to the benthos. The biodeposition method is used to quantify feeding variables in suspension-feeding bivalves in a natural setting and represents a more realistic proxy than laboratory experiments. This method, however, relies upon a stable platform to satisfy the requirements that water flow rates supplied to the shellfish remain constant and the bivalves are undisturbed. A flow-through device and process for using the biodeposition method to quantify the feeding of bivalve mollusks were modified from a land-based format for shipboard use by building a two-dimensional gimbal table around the device. Planimeter data reveal a minimal pitch and yaw of the chambers containing the test shellfish despite boat motion, the flow rates within the chambers remain constant, and operators are able to collect the biodeposits (feces and pseudofeces) with sufficient consistency to obtain accurate measurements of the bivalve clearance, filtration, selection, ingestion, rejection, and absorption at offshore shellfish aquaculture sites.

A flow-through device for using the biodeposition method to quantify filtration and feeding behavior of bivalve mollusks was modified for shipboard use. A two-dimensional gimbal table built around the device isolates the apparatus from boat motion, thereby allowing the accurate quantification of bivalve filtration variables at offshore shellfish aquaculture sites.

As shellfish aquaculture moves from coastal embayments and estuaries to offshore locations, the need to quantify ecosystem interactions of farmed bivalves ...

Development and Testing of Species-specific Quantitative PCR Assays for Environmental DNA Applications

New, non-invasive methods for detecting and monitoring species presence are being developed to aid in fisheries and wildlife conservation management. The use of environmental DNA (eDNA) samples for detecting macrobiota is one such group of methods that is rapidly becoming popular and being implemented in national management programs. Here we focus on the development of species-specific targeted assays for probe-based quantitative PCR (qPCR) applications. Using probe-based qPCR offers greater specificity than is possible with primers alone. Furthermore, the ability to quantify the amount of DNA in a sample can be useful in our understanding of the ecology of eDNA and the interpretation of eDNA detection patterns in the field. Careful consideration is needed in the development and testing of these assays to ensure the sensitivity and specificity of detecting the target species from an environmental sample. In this protocol we will delineate the steps needed to design and test probe-based assays for the detection of a target species; including creation of sequence databases, assay design, assay selection and optimization, testing assay performance, and field validation. Following these steps will help achieve an efficient, sensitive, and specific assay that can be used with confidence. We demonstrate this process with our assay designed for populations of the mucket (Actinonaias ligamentina), a freshwater mussel species found in the Clinch River, USA.

Environmental DNA assays require rigorous design, testing, optimization and validation before the collection of field data can begin. Here, we present a protocol to take users through each step of designing a species-specific, probe-based qPCR assay for the detection and quantification of a target species DNA from environmental samples.

New, non-invasive methods for detecting and monitoring species presence are being developed to aid in fisheries and wildlife conservation management. The ...

Hellgrammite as a Non-Conventional Biomonitor for Surface Water and Environmental Health Assessment

The larvae of dobsonflies from the genus Corydalus, commonly known as Hellgrammites, are characterized by their notable size, extensive range of occurrence, and extended period of immaturity, which can last up to one year. Hellgrammites are clearly known to exhibit sensitivity to pollution and habitat structure impacts. Given these unique features, using Corydalus texanus larvae is highly suitable as reliable biomonitoring agents to assess the ecological integrity of aquatic ecosystems. This protocol aims to provide the necessary tools for C. texanus assessment and demonstrate their efficacy through a case study. Research findings have practical implications, indicating that C. texanus larvae exhibit early-warning responses to mining pollution, bioaccumulating high amounts of heavy metals such as Zn, Fe, and Al. The presence or absence of C. texanus populations may serve as a helpful indicator for identifying potential issues related to ecosystem health. The unconventional approach has shown early warnings of pollution in mining-impacted sites, highlighting the need for timely action to protect the environment. Given their unique traits, the use of C. texanus larvae is highly suggested as a reliable non-conventional bioindicator.

This protocol describes the use of biomarkers for the early detection of deleterious impacts in aquatic ecosystems. The biomarkers are closely related to sentinel traits, and their changes aid in detecting early-warning damages.

The larvae of dobsonflies from the genus Corydalus, commonly known as Hellgrammites, are characterized by their notable size, extensive range of ...

Standardizing a Non-Lethal Method for Characterizing the Reproductive Status and Larval Development of Freshwater Mussels (Bivalvia: Unionida)

Actively monitoring the timing, development, and reproductive patterns of endangered species is critical when managing for population recovery. Freshwater mussels are among the most imperiled organisms in the world, but information about early larval (glochidial) development and brooding periods is still lacking for many species. Previous studies have focused on the complex life history stage when female mussels are ready to parasitize host fish, but few studies have focused on the brooding period and timing of larval development. The protocol described here allows researchers to non-lethally evaluate the state of gravidity for female mussels. The results of this study show that this method does not affect a female mussel&rsquo;s ability to stay gravid or become gravid again after sampling has been performed. The advantage of this method may permit its use on federally threatened or endangered species or other populations of high conservation concern. This protocol can be adapted for use on both preserved or live individuals and was tested on a variety of mussel species. The database provided is a repository for a breadth of information on timing of reproductive habits and will facilitate future freshwater mussel research, conservation, and recovery efforts.

Standardizing a Non-Lethal Method for Characterizing the Reproductive Status and Larval Development of Freshwater Mussels Bivalvia: Unionida

Freshwater mussel conservation is dependent on monitoring reproductive patterns and processes of species. This study standardizes a non-lethal protocol for sampling gill contents, characterizing larval development, and providing a digital repository for data collected. This protocol-database package will be an important tool for mussel researchers in recovery of imperiled species.

Actively monitoring the timing, development, and reproductive patterns of endangered species is critical when managing for population recovery. Freshwater ...

Biology

A Toxicological and Ecotoxicological Assay Based on Mussel (Mytilus galloprovincialis) Hemocytes Motility

Hemocytes are the circulating immune-competent cells in bivalve mollusks and play a key role in several important functions of cell-mediated innate immunity. During the early stages of the immune response, hemocytes actively migrate to the site of infection. This inherent motility is a fundamental characteristic of these cells. It represents a key cellular function that integrates multiple processes, such as cell adhesion, cell signaling, cytoskeletal dynamics, and changes in cell volume. Therefore, alterations in cell motility following exposure to drugs or pollutants can serve as a useful toxicological endpoint. Despite the fundamental role of cell motility in cellular physiology, it has been poorly investigated from a toxicological perspective. This work proposes a novel in vitro method for the rapid and sensitive assessment of the toxicity and ecotoxicity of pollutants, based on evaluating the hemocyte motility of Mytilus galloprovincialis. We developed a cell motility assay on hemocytes adhering to the bottom of a 96-well polystyrene microplate. Following exposure to increasing concentrations of drugs, cell trajectories, and velocities were quantified by cell tracking under time-lapse microscopy, allowing us to measure the effects on hemocyte motility. Due to the ease of hemocyte collection from the animals in a relatively non-invasive manner, the proposed method offers an alternative test for screening the effects and mechanisms of action of pollutants and drugs. It aligns with the 3Rs (Replacement, Reduction, and Refinement) criteria, addressing ethical concerns and contributing to the reduction of vertebrate in vivo animal testing.

The article proposes a novel in vitro method for the rapid and sensitive assessment of the toxicity and ecotoxicity of pollutants, based on the motility of Mytilus galloprovincialis hemocytes. The method aims to contribute to the development of more ethical and sensitive toxicological and ecotoxicological exposure tests.

Hemocytes are the circulating immune-competent cells in bivalve mollusks and play a key role in several important functions of cell-mediated innate ...

Bioindication Testing of Stream Environment Suitability for Young Freshwater Pearl Mussels Using In Situ Exposure Methods

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