Funding for composing and publishing this paper was provided through multiple grants and contracts to the Chironomidae Research Group (L. C. Ferrington, Jr., PI) in the Department of Entomology at the University of Minnesota. &#160;Thanks to Nathan Roberts for sharing fieldwork photographs used as figures in the video associated with this manuscript.

1. Preparation of Field Collection Supplies
<ol>
	<li>Determine the number of SFPE samples that should be collected based on the study design and acquire one sample jar (e.g., 60 ml) for each sample.</li>
	<li>Prepare two date and locality labels for each sample jar. Place one on the inside and affix the other to the outside of the jar. Ensure that each date and locality label includes the following information:&#160;&#160;country, state, county, city, water body, GPS coordinates, date, and name of person(s) collecting the sample.</li>
	<li>Gather other specific materials and equipment (see Table of Specific Materials/Equipment).</li>
</ol>
2. Field Collection
<ol>
	<li>Hold a larval tray in one hand and a sieve in the other. Dip the larval tray into the water where SFPE accumulate (e.g., foam accumulations, snags, emergent vegetation, debris, back eddies, and along bank edges) (Figure 2A), allow water, exuviae, and debris to enter the larval tray, and pour this material through the sieve. If sampling in a lotic system, begin at the downstream end of the sample reach and work upstream (Figure 2B). If sampling in a lentic system, begin at the downwind shoreline.&#160;
	<ol>
		<li>Repeat step 2.1 for 10 min (or as otherwise defined for a specific sampling regime)&#160;within each pre-defined sample reach (typically 100-200 m for samples collected from streams, but dependent on the overall area of the aquatic monitoring site); move between SFPE accumulation areas as appropriate.</li>
	</ol>
	</li>
	<li>Concentrate debris in one area of the sieve using a squirt bottle filled with water from the sample site and carefully transfer SFPE sample to pre-labeled sample jar with the aid of forceps and a stream of ethanol from a squirt bottle. Fill sample jar with ethanol.</li>
	<li>Repeat steps 2.1 to 2.2 for all samples.</li>
</ol>
3. Sample Picking
NOTE: The rest of this protocol pertains to a 300 SFPE subsample and may need to be modified for other subsample sizes. See Bouchard and Ferrington&#39;s9 subsampling and sampling frequency guidelines for tailoring SFPE methods to meet study-specific goals and resources.
<ol>
	<li>Allocate a 1-dram vial for each SFPE sample; prepare a date and &#160;locality label to place inside each vial and fill the vial &#190; full with ethanol.</li>
	<li>Remove lid from the corresponding sample jar and check for attached pupal exuviae. Gently rinse contents off the lid onto a Petri dish using a squirt bottle filled with ethanol. Locate and remove label from the inside of the sample jar using forceps and gently rinse contents off the label onto the Petri dish. Set label aside.</li>
	<li>Transfer the contents of the sample jar into a&#160;larval tray, rinsing with ethanol to ensure no SFPE remain in the sample jar. Transfer a portion of the pupal exuviae, residue, and ethanol from the tray to the Petri dish. Ensure that the sample is covered in ethanol.&#160;</li>
	<li>Place the Petri dish under a stereo microscope. Systematically scan the contents of the Petri dish for pupal exuviae. Pick all pupal exuviae from the dish using forceps and place into the vial. Do not pick specimens that are broken (i.e., do not have at least half of the cephalothorax and abdomen), dried, or compressed to avoid later identification problems. 
	NOTE: Identification to species often requires that&#160;the entire specimen is present, though&#160;in some cases,&#160;genus-level identification may be possible with partial specimens.
	<ol>
		<li>Swirl dish and scan for additional pupal exuviae, including any that could be stuck to sides of the dish, as well as, any small and translucent specimens that may not have be detected initially. Repeat until two consecutive scans reveal no additional pupal exuviae.</li>
	</ol>
	</li>
	<li>Repeat steps 3.3 and 3.4 until all or 300 pupal exuviae have been picked.&#160;When 300 pupal exuviae have been picked, return the residue from the Petri dish to the larval tray and rinse the Petri dish with ethanol. Then, transfer the residue from the larval tray to the empty sample jar, add the date and locality label, and put the lid on&#160; the jar. Retain or dispose of residue according to project-specific protocols.</li>
</ol>
4. Sample Sorting
<ol>
	<li>Pour all picked pupal exuviae from the labeled vial into a Petri dish filled with enough ethanol to just cover specimens.</li>
	<li>Under a stereo microscope, separate specimens into distinct morphological groups (i.e., morphotaxa) and place each morphotaxon into separately&#160;a labeled vials filled 3/4th full with ethanol.
	<ol>
		<li>Utilize external morphological characteristics to separate chironomid morphotaxa. For example, from the cephalothorax, use differences in the presence, size, shape, and coloration of the cephalic tubercles, frontal warts, frontal setae, and thoracic horn. From the abdomen, use spines, hookrows, shagreen, setae, and spurs of the abdominal segments, in addition to the anal lobes&#160;for morphotaxa separation (Figure 4A). See Ferrington, et al. 5,S&#230;ther 26,Pinder and Reiss 27 for additional descriptions and figures of morphological characteristics.</li>
		<li>Use additional ethanol if specimens begin to dry.</li>
	</ol>
	</li>
</ol>
5. Slide Mounting
<ol>
	<li>Fill one well of a multi-well plate for each morphotaxon with 95% ethanol.
	<ol>
		<li>Place multiple representations (e.g., 25% of total) of each morphotaxon to be slide mounted into individual wells of the plate. Allow specimens to sit in well for at least 10 min&#160;to dehydrate&#160;sufficiently.</li>
	</ol>
	</li>
	<li>Label slides with appropriate site, collection, and identification information (Figure 3).</li>
	<li>Place slide on the stereo microscope. 
	NOTE: A template of the slide taped to the stage is useful for consistent placement.</li>
	<li>Place a drop of Euparal on the slide; spread the Euparal so that it approximates the size of the coverslip. Use proper ventilation when working with Euparal. 
	NOTE: Use proper ventilation when working with Euparal.</li>
	<li>Embed a representative from the first morphotaxon into the Euparal using forceps. 
	NOTE: To void excess ethanol from the specimen, using a forceps, gently tap specimen on laboratory wipes &#160;prior to embedding it in Euparal.</li>
	<li>Separate the cephalothorax from the abdomen using fine-tipped forceps and/or dissection probes (Figure 4A).
	<ol>
		<li>Split the cephalothorax along the ecdysial suture (Figure 4B) and open the cephalothorax so that the suture edges are on opposite sides (Figure 4C).</li>
		<li>Orient the cephalothorax so that the ventral side is facing up (Figure 4C).</li>
		<li>Position the abdomen dorsal side up; place immediately below the cephalothorax (Figure 4C).</li>
	</ol>
	</li>
	<li>Place a coverslip on the specimen. Hold coverslip at an angle, with one edge touching the slide, and then slowly lower and drop the coverslip to reduce air bubble formation. Press lightly on the coverslip to flatten the specimen.</li>
	<li>Repeat steps 5.3 through 5.7 for all dehydrated&#160;specimens.</li>
</ol>
6. Genus Identification
<ol>
	<li>Determine genus of slide-mounted specimens using a compound microscope. Identify specimens to genus using keys and diagnoses in Wiederholm 28 and Ferrington, et al.5. If needed, confirm family-level identification using Ferrington, et al.5. &#160;&#160;NOTE: There have been numerous generic descriptions and revisions since Wiederholm28 and Ferrington, et al.5, therefore, these keys and diagnoses are incomplete and need to be supplemented with primary literature.</li>
</ol>

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The authors declare that they have no competing financial interests.

The most critical steps for successful SFPE sample collection, picking, sorting, slide mounting, and identification are: (1) locating areas of high SFPE accumulation within the study area during field collection (Figure 2A); (2) slowly scanning the contents of the Petri dish for detection of all&#160;SFPE during sample picking; (3) developing the necessary manual dexterity to dissect the cephalothorax from the abdomen during slide mounting (Figure 4A); and (4) recognizing key morphologic...

Biological monitoring programs, which use living organisms to evaluate environmental health, are often used to assess water quality or monitor success of ecosystem restoration programs. Rapid bioassessment protocols (RBP) using benthic macroinvertebrate assemblages have been popular among state water resource agencies since 19891. Traditional methods of sampling benthic macroinvertebrates for RBPs, such as the dip-net, Surber sampler, and Hess sampler2, can be time-consuming, expensive, and may only measure assemblages from a particular microhabitat3. An efficient, alternative RBP for generating biological information about a particular water body involves collection of Chironomidae surface-floating pupal exuviae (SFPE)3.
The Chironomidae (Insecta: Diptera), commonly known as non-biting midges, are holometabolous flies that typically occur in aquatic environments before emerging as adults&#160;on the water&#8217;s surface. The chironomid family is species-rich, with approximately 5,000 species described worldwide; however, as many as 20,000 species are estimated to exist4. Chironomids are useful in documenting water and habitat quality in many aquatic ecosystems because of their high diversity and variable pollution tolerance levels5. Furthermore, they are often the most abundant and widespread benthic macroinvertebrates in aquatic systems, typically accounting for 50% or more of the species in the community5,6. Following emergence of the terrestrial adult, the pupal exuviae (cast pupal skin) remains floating on the water&#8217;s surface (Figure 1). Pupal exuviae accumulate along banks or behind obstructions through the action of wind or water current and can be easily and rapidly collected to give a comprehensive sample of chironomid species that have emerged during the previous 24-48 hr7.
The relative abundance and taxonomic composition of collected SFPE reflects water quality, considering that some species are very pollution tolerant, while others are quite sensitive5. The SFPE method has many advantages over traditional larval chironomid sampling techniques including: (1) minimal, if any, habitat disturbance occurs at a sampling area; (2) samples do not focus on collecting living organisms, but rather the non-living skin, so the trajectory of community dynamics is not affected; (3) identification to genus, and often species, is relatively easy given appropriate keys and descriptions3; (4) collecting, processing, and identifying samples is efficient and economical in comparison to traditional sampling methods3,8,9; (5) accumulated exuviae represent taxa that have originated from a wide range of microhabitats10; (6) the method is applicable in nearly all aquatic environments, including streams and rivers, estuaries, lakes, ponds, rock pools, and wetlands; and (7) SFPE maybe be a more sensitive indicator of ecosystem health since they represent individuals that have completed all immature stages and successfully emerged as&#160;adults11.
The SFPE method is not a new approach for gathering information about chironomid communities. Use of SFPE was first suggested by Thienemann12 in the early 1900s. A variety of studies have used SFPE for taxonomic surveys (e.g.,&#160;13-15), biodiversity and ecological studies (e.g. 7,16-19), and biological assessments (e.g.,20-22). Additionally, some studies have addressed different aspects of sample design, sample size, and number of sample events required for achieving various detection levels of species or genera (e.g.,8,9,23). These studies indicate that relatively high percentages of species or genera can be detected with moderate effort or expense associated with sample processing. For example, Anderson and Ferrington8 determined that based on a 100-count subsample, &#160;1/3rd less time was required&#160;to pick SFPE samples compared to dip-net samples. Another study determined that 3-4 SFPE samples could be sorted and identified for every dip-net sample and that SFPE samples were more efficient than dip-net samples at detecting species as species richness increased3. For example, at sites with species richness values of 15-16 species, the average dip-net efficiency was 45.7%, while SFPE samples were 97.8% efficient3.
Importantly, the SFPE method has been standardized in the European Union24 (known as chironomid pupal exuviae technique (CPET)) and North America25 for ecological assessment, but the method has not been described in detail. One application of the SFPE methodology was described by Ferrington, et al.3; however, the primary focus of that study was to evaluate the efficiency, efficacy, and economy of the SFPE method. The purpose of this work is to describe all steps of the SFPE method in detail, including sample collection, laboratory processing, slide mounting, and genus identification. The target audience includes graduate students, researchers, and professionals interested in expanding traditional water quality monitoring efforts into their studies.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Ethanol</td>
			<td>Fisher Scientific</td>
			<td>S25309B&#160;</td>
			<td>70-95%</td>
		</tr>
		<tr>
			<td>Plastic wash bottles</td>
			<td>Fisher Scientific</td>
			<td>0340923B</td>
			<td></td>
		</tr>
		<tr>
			<td>Sample jar</td>
			<td>Fisher Scientific</td>
			<td>0333510B</td>
			<td>Glass or plastic, 60-mL recommended</td>
		</tr>
		<tr>
			<td>Testing sieve</td>
			<td>Advantech</td>
			<td>120SS12F</td>
			<td>125-micron mesh size</td>
		</tr>
		<tr>
			<td>Larval tray</td>
			<td>BioQuip</td>
			<td>5524</td>
			<td>White</td>
		</tr>
		<tr>
			<td>Stereo microscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass shell vials</td>
			<td>Fisher Scientific</td>
			<td>0333926B</td>
			<td>1-dram size</td>
		</tr>
		<tr>
			<td>Plastic dropper</td>
			<td>Thermo Scientific</td>
			<td>1371110</td>
			<td>30 to 35 drops/mL</td>
		</tr>
		<tr>
			<td>Fine forceps</td>
			<td>BioQuip</td>
			<td>4524</td>
			<td>#5</td>
		</tr>
		<tr>
			<td>Petri dish</td>
			<td>Carolina</td>
			<td>741158</td>
			<td>Glass or plastic</td>
		</tr>
		<tr>
			<td>Multi-well plate</td>
			<td>Thermo Scientific</td>
			<td align="right">144530</td>
			<td>Glass or plastic</td>
		</tr>
		<tr>
			<td>Glass microslides</td>
			<td>Thermo Scientific</td>
			<td>3010002</td>
			<td>3 x 1 in.</td>
		</tr>
		<tr>
			<td>Glass cover slips</td>
			<td>Thermo Scientific</td>
			<td>12-519-21G</td>
			<td>Circular or square</td>
		</tr>
		<tr>
			<td>Euparal mounting medium&#160;</td>
			<td>BioQuip</td>
			<td>6372B</td>
			<td></td>
		</tr>
		<tr>
			<td>Pigma pen</td>
			<td>BioQuip</td>
			<td>1154F</td>
			<td>Black</td>
		</tr>
		<tr>
			<td>Probe</td>
			<td>BioQuip</td>
			<td>4751</td>
			<td></td>
		</tr>
		<tr>
			<td>Kimwipes</td>
			<td>Kimberly-Clark Professional&#8482;</td>
			<td align="right">34120</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Figure 1 illustrates the chironomid life cycle; immature stages (egg, larva, pupa) typically take place in, or closely associated with, an aquatic environment. Upon completion of the larval life stage, the larva constructs a tube-like shelter and attaches itself with silken secretions to the surrounding substrate and pupation occurs. Once the developing adult has matured, the pupa frees itself and swims to the surface of the water where the adult can emerge from the pupal exuviae. The exuviae fills with ...

Watch this Scientific Journal Video about Use of Chironomidae Diptera Surface-Floating Pupal Exuviae as a Rapid Bioassessment Protocol for Water Bodies at JoVE.com

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.

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. ...

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11.6K ビュー.  University of Minnesota. 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.

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

Laboratory-determined Phosphorus Flux from Lake Sediments as a Measure of Internal Phosphorus Loading

Eutrophication is a water quality issue in lakes worldwide, and there is a critical need to identify and control nutrient sources. Internal phosphorus (P) loading from lake sediments can account for a substantial portion of the total P load in eutrophic, and some mesotrophic, lakes. Laboratory determination of P release rates from sediment cores is one approach for determining the role of internal P loading and guiding management decisions. Two principal alternatives to experimental determination of sediment P release exist for estimating internal load: in situ measurements of changes in hypolimnetic P over time and P mass balance. The experimental approach using laboratory-based sediment incubations to quantify internal P load is a direct method, making it a valuable tool for lake management and restoration.
Laboratory incubations of sediment cores can help determine the relative importance of internal vs. external P loads, as well as be used to answer a variety of lake management and research questions. We illustrate the use of sediment core incubations to assess the effectiveness of an aluminum sulfate (alum) treatment for reducing sediment P release. Other research questions that can be investigated using this approach include the effects of sediment resuspension and bioturbation on P release.
The approach also has limitations. Assumptions must be made with respect to: extrapolating results from sediment cores to the entire lake; deciding over what time periods to measure nutrient release; and addressing possible core tube artifacts. A comprehensive dissolved oxygen monitoring strategy to assess temporal and spatial redox status in the lake provides greater confidence in annual P loads estimated from sediment core incubations.

Lake eutrophication is a water quality issue worldwide, making the need to identify and control nutrient sources critical. Laboratory determination of phosphorus release rates from sediment cores is a valuable approach for determining the role of internal phosphorus loading and guiding management decisions.

Eutrophication is a water quality issue in lakes worldwide, and there is a critical need to identify and control nutrient sources. Internal phosphorus (P) ...

The Use of an Automated System (GreenFeed) to Monitor Enteric Methane and Carbon Dioxide Emissions from Ruminant Animals

Ruminant animals (domesticated or wild) emit methane (CH4) through enteric fermentation in their digestive tract and from decomposition of manure during storage. These processes are the major sources of greenhouse gas (GHG) emissions from animal production systems. Techniques for measuring enteric CH4 vary from direct measurements (respiration chambers, which are highly accurate, but with limited applicability) to various indirect methods (sniffers, laser technology, which are practical, but with variable accuracy). The sulfur hexafluoride (SF6)&#160;tracer gas method is commonly used to measure enteric CH4 production by animal scientists and more recently, application of an Automated Head-Chamber System (AHCS) (GreenFeed, C-Lock, Inc., Rapid City, SD), which is the focus of this experiment, has been growing. AHCS is an automated system to monitor CH4 and carbon dioxide (CO2) mass fluxes from the breath of ruminant animals. In a typical AHCS operation, small quantities of baiting feed are dispensed to individual animals to lure them to AHCS multiple times daily. As the animal visits AHCS, a fan system pulls air past the animal&#8217;s muzzle into an intake manifold, and through an air collection pipe where continuous airflow rates are measured. A sub-sample of air is pumped out of the pipe into non-dispersive infra-red sensors for continuous measurement of CH4 and CO2 concentrations. Field comparisons of AHCS to respiration chambers or SF6 have demonstrated that AHCS produces repeatable and accurate CH4 emission results, provided that animal visits to AHCS are sufficient so emission estimates are representative of the diurnal rhythm of rumen gas production. Here, we demonstrate the use of AHCS to measure CO2 and CH4 fluxes from dairy cows given a control diet or a diet supplemented with technical-grade cashew nut shell liquid.

The Use of an Automated System GreenFeed to Monitor Enteric Methane and Carbon Dioxide Emissions from Ruminant Animals

Accuracy and precision of the techniques used to measure methane emissions from ruminant animals are critically important for the success of greenhouse gas mitigation efforts. This manuscript describes the principles and operation of an automated system to monitor methane and carbon dioxide mass fluxes from the breath of ruminant animals.

Ruminant animals (domesticated or wild) emit methane (CH4) through enteric fermentation in their digestive tract and from decomposition of manure during ...

A CO2 Concentration Gradient Facility for Testing CO2 Enrichment and Soil Effects on Grassland Ecosystem Function

Continuing increases in atmospheric carbon dioxide concentrations (CA) mandate techniques for examining impacts on terrestrial ecosystems. Most experiments examine only two or a few levels of CA concentration and a single soil type, but if CA can be varied as a gradient from subambient to superambient concentrations on multiple soils, we can discern whether past ecosystem responses may continue linearly in the future and whether responses may vary across the landscape. The Lysimeter Carbon Dioxide Gradient Facility applies a 250 to 500 &#181;l L-1 CA gradient to Blackland prairie plant communities established on lysimeters containing clay, silty clay, and sandy soils. The gradient is created as photosynthesis by vegetation enclosed in in temperature-controlled chambers progressively depletes carbon dioxide from air flowing directionally through the chambers. Maintaining proper air flow rate, adequate photosynthetic capacity, and temperature control are critical to overcome the main limitations of the system, which are declining photosynthetic rates and increased water stress during summer. The facility is an economical alternative to other techniques of CA enrichment, successfully discerns the shape of ecosystem responses to subambient to superambient CA enrichment, and can be adapted to test for interactions of carbon dioxide with other greenhouse gases such as methane or ozone.

A CO2 Concentration Gradient Facility for Testing CO2 Enrichment and Soil Effects on Grassland Ecosystem Function

The Lysimeter Carbon Dioxide Gradient Facility creates a 250 to 500 &#181;l L-1 linear carbon dioxide gradient in temperature-controlled chambers housing grassland plant communities on clay, silty clay, and sandy soil monoliths. The facility is used to determine how past and future carbon dioxide levels affect grassland carbon cycling.

Continuing increases in atmospheric carbon dioxide concentrations (CA) mandate techniques for examining impacts on terrestrial ecosystems. Most ...

Preparation and Testing of Impedance-based Fluidic Biochips with RTgill-W1 Cells for Rapid Evaluation of Drinking Water Samples for Toxicity

This manuscript describes how to prepare fluidic biochips with Rainbow trout gill epithelial (RTgill-W1) cells for use in a field portable water toxicity sensor. A monolayer of RTgill-W1 cells forms on the sensing electrodes enclosed within the biochips. The biochips are then used for testing in a field portable electric cell-substrate impedance sensing (ECIS) device designed for rapid toxicity testing of drinking water. The manuscript further describes how to run a toxicity test using the prepared biochips. A control water sample and the test water sample are mixed with pre-measured powdered media and injected into separate channels of the biochip. Impedance readings from the sensing electrodes in each of the biochip channels are measured and compared by an automated statistical software program. The screen on the ECIS instrument will indicate either "Contamination Detected" or "No Contamination Detected" within an hour of sample injection. Advantages are ease of use and rapid response to a broad spectrum of inorganic and organic chemicals at concentrations that are relevant to human health concerns, as well as the long-term stability of stored biochips in a ready state for testing. Limitations are the requirement for cold storage of the biochips and limited sensitivity to cholinesterase-inhibiting pesticides. Applications for this toxicity detector are for rapid field-portable testing of drinking water supplies by Army Preventative Medicine personnel or for use at municipal water treatment facilities.

This manuscript describes how to prepare fluidic biochips with Rainbow trout gill epithelial cells for use in a field portable electric cell-substrate impedance sensor. The protocol for running a rapid drinking water toxicity test with the sensor is also described.

This manuscript describes how to prepare fluidic biochips with Rainbow trout gill epithelial (RTgill-W1) cells for use in a field portable water toxicity ...

Protocol for Measuring the Thermal Properties of a Supercooled Synthetic Sand-water-gas-methane Hydrate Sample

Methane hydrates (MHs) are present in large amounts in the ocean floor and permafrost regions. Methane and hydrogen hydrates are being studied as future energy resources and energy storage media. To develop a method for gas production from natural MH-bearing sediments and hydrate-based technologies, it is imperative to understand the thermal properties of gas hydrates.
The thermal properties' measurements of samples comprising sand, water, methane, and MH are difficult because the melting heat of MH may affect the measurements. To solve this problem, we performed thermal properties' measurements at supercooled conditions during MH formation. The measurement protocol, calculation method of the saturation change, and tips for thermal constants' analysis of the sample using transient plane source techniques are described here.
The effect of the formation heat of MH on measurement is very small because the gas hydrate formation rate is very slow. This measurement method can be applied to the thermal properties of the gas hydrate-water-guest gas system, which contains hydrogen, CO2, and ozone hydrates, because the characteristic low formation rate of gas hydrate is not unique to MH. The key point of this method is the low rate of phase transition of the target material. Hence, this method may be applied to other materials having low phase-transition rates.

We present a protocol for measuring the thermal properties of synthetic hydrate-bearing sediment samples comprising sand, water, methane, and methane hydrate.

Methane hydrates (MHs) are present in large amounts in the ocean floor and permafrost regions. Methane and hydrogen hydrates are being studied as future ...

Soil Lysimeter Excavation for Coupled Hydrological, Geochemical, and Microbiological Investigations

Studying co-evolution of hydrological and biogeochemical processes in the subsurface of natural landscapes can enhance the understanding of coupled Earth-system processes. Such knowledge is imperative in improving predictions of hydro-biogeochemical cycles, especially under climate change scenarios. We present an experimental method, designed to capture sub-surface heterogeneity of an initially homogeneous soil system. This method is based on destructive sampling of a soil lysimeter designed to simulate a small-scale hillslope. A weighing lysimeter of one cubic meter capacity was divided into sections (voxels) and was excavated layer-by-layer, with sub samples being collected from each voxel. The excavation procedure was aimed at detecting the incipient heterogeneity of the system by focusing on the spatial assessment of hydrological, geochemical, and microbiological properties of the soil. Representative results of a few physicochemical variables tested show the development of heterogeneity. Additional work to test interactions between hydrological, geochemical, and microbiological signatures is planned to interpret the observed patterns. Our study also demonstrates the possibility of carrying out similar excavations in order to observe and quantify different aspects of soil-development under varying environmental conditions and scale.

This study presents an excavation method for investigating subsurface hydrological, geochemical, and microbiological heterogeneity of a soil lysimeter. The lysimeter simulates an artificial hillslope which was initially under homogeneous condition and had been subjected to approximately 5,000 mm of water over eight cycles of irrigation in an 18-month period.

Studying co-evolution of hydrological and biogeochemical processes in the subsurface of natural landscapes can enhance the understanding of coupled ...

Use of a Filter Cartridge for Filtration of Water Samples and Extraction of Environmental DNA

Recent studies demonstrated the use of environmental DNA (eDNA) from fishes to be appropriate as a non-invasive monitoring tool. Most of these studies employed disk fiber filters to collect eDNA from water samples, although a number of microbial studies in aquatic environments have employed filter cartridges, because the cartridge has the advantage of accommodating large water volumes and of overall ease of use. Here we provide a protocol for filtration of water samples using the filter cartridge and extraction of eDNA from the filter without having to cut open the housing. The main portions of this protocol consists of 1) filtration of water samples (water volumes &#8804;4 L or &gt;4 L); (2) extraction of DNA on the filter using a roller shaker placed in a preheated incubator; and (3) purification of DNA using a commercial kit. With the use of this and previously-used protocols, we perform metabarcoding analysis of eDNA taken from a huge aquarium tank (7,500 m3) with known species composition, and show the number of detected species per library from the two protocols as the representative results. This protocol has been developed for metabarcoding eDNA from fishes, but is also applicable to eDNA from other organisms.

We describe a protocol for filtration of water samples with a filter cartridge and extraction of environmental DNA (eDNA) without having to cut open the housing to remove the filter. This protocol is developed for metabarcoding eDNA from fishes, but is also applicable to eDNA from other organisms.

Recent studies demonstrated the use of environmental DNA (eDNA) from fishes to be appropriate as a non-invasive monitoring tool. Most of these studies ...

Protocol for Microplastics Sampling on the Sea Surface and Sample Analysis

Microplastic pollution in the marine environment is a scientific topic that has received increasing attention over the last decade. The majority of scientific publications address microplastic pollution of the sea surface. The protocol below describes the methodology for sampling, sample preparation, separation and chemical identification of microplastic particles. A manta net fixed on an &#187;A frame&#171; attached to the side of the vessel was used for sampling. Microplastic particles caught in the cod end of the net were separated from samples by visual identification and use of stereomicroscopes. Particles were analyzed for their size using an image analysis program and for their chemical structure using ATR-FTIR and micro FTIR spectroscopy. The described protocol is in line with recommendations for microplastics monitoring published by the Marine Strategy Framework Directive (MSFD) Technical Subgroup on Marine Litter. This written protocol with video guide will support the work of researchers that deal with microplastics monitoring all over the world.

The protocol below describes the methodology for: microplastics sampling on the sea surface, separation of microplastic and chemical identification of particles. This protocol is in line with the recommendations for microplastics monitoring published by the MSFD Technical Subgroup on Marine Litter.

Microplastic pollution in the marine environment is a scientific topic that has received increasing attention over the last decade. The majority of ...

Measuring and Mapping Patterns of Soil Erosion and Deposition Related to Soil Carbonate Concentrations Under Agricultural Management

Spatial patterns of soil erosion and deposition can be inferred from differences in ground elevation mapped at appropriate time increments. Such changes in elevation are related to changes in near-surface soil carbonate (CaCO3) profiles. The objective is to describe a simple conceptual model and detailed protocol for repeatable field and laboratory measurements of these quantities. Here, accurate elevation is measured using a ground-based differential global positioning system (GPS); other data acquisition methods could be applied to the same basic method. Soil samples are collected from prescribed depth intervals and analyzed in the lab using an efficient and precise modified pressure-calcimeter method for quantitative analysis of inorganic carbon concentration. Standard statistical methods are applied to point data, and representative results show significant correlations between changes in soil surface layer CaCO3 and changes in elevation consistent with the conceptual model; CaCO3 generally decreased in depositional areas and increased in erosional areas. Maps are derived from point measurements of elevation and soil CaCO3 to aid analyses. A map of erosional and depositional patterns at the study site, a rain-fed winter wheat field cropped in alternating wheat-fallow strips, shows the interacting effects of water and wind erosion affected by management and topography. Alternative sampling methods and depth intervals are discussed and recommended for future work relating soil erosion and deposition to soil CaCO3.

Spatial patterns of soil erosion and deposition can be inferred from differences in ground elevation mapped at appropriate time increments. Such changes in elevation are related to changes in near-surface soil carbonates. Repeatable methods for field and laboratory measurements of these quantities and data analysis methods are described here.

Spatial patterns of soil erosion and deposition can be inferred from differences in ground elevation mapped at appropriate time increments. Such changes ...

A Simple Planting Technique for Re-establishing Trees Where Frequent Inundation Occurs

Many of the Everglades tree islands have lost elevation over the past century and most of their trees have died such that they are now covered with herbaceous plants. This protocol describes a simple, cost-effective tree planting technique needed for restoring degraded Everglade tree islands. The design is patterned after a natural Everglades process that creates floating peat islands, which allows tree survival and growth in flooded conditions and often leads to the development of tree islands. Commercially available peat bags were used as the medium for the growth and establishment of potted native tree saplings. The pop-up configuration floated initially and provided additional elevation to minimize inundation, with a single native tree species sapling and a single tree fertilizer spike. During a 3 year study involving 105 pop-ups, most plants survived (80%) and many thrived. Determining whether this technique can establish trees on a degraded tree island will require longer studies and extensive field tests.

This protocol describes a simple, cost-effective tree planting technique for restoring degraded Everglades tree islands that experience inundation. The design creates an island (pop-up) which floats initially and adds elevation to promote tree survival and growth under flooded conditions.

Many of the Everglades tree islands have lost elevation over the past century and most of their trees have died such that they are now covered with ...