The authors thank Drs. Bobby J. Presley, Robert Tayloy, Paul Boothe, Mr. Bryan Brattin, and Mr. Mike Metcalf for their assistance during the laborious field sampling and lab work for the practical development and application of "clean techniques".

1. Sampler Preparation
<ol>
	<li>Sampler
	<ol>
		<li>Assemblage of sampler
		<ol>
			<li>Connect a 4 m long fluorinated ethylene propylene (FEP) tubing (I.D. 0.635 cm, O.D. 0.95 cm or similar) to a 1.5-m chemically resistant silicone pumping tube (O.D. 0.635 cm).</li>
			<li>Insert a polypropylene Y-connector into the pumping tube, and connect a 50-cm pumping tube to one outlet, and a 0.45 &#956;m capsule filter (by a 20-cm pumping tube) to the other.</li>
			<li>Assemble the tubings in a clean room (bench) after they are cleaned (see below), and store the assemblage in two layers of polyethylene bags.</li>
		</ol>
		</li>
		<li>Sampler cleaning
		<ol>
			<li>Fill (by attaching the 1.5-m pumping tube onto a peristaltic pump) the tubing set with lab detergent solution and soak it for 24 hr. Flush the tubing set with de-ionized water, then fill it with 10% (v/v) HCl (Reagent Grade) and soak for 48 hr.</li>
			<li>Thoroughly flush the tubing set with de-ionized water several times, and store the assemblage in plastic bags. Clean the Y-connectors and short pumping tubes by soaking them in a 50% (v/v) HNO3 (Reagent Grade) solution for 24 hr.</li>
		</ol>
		</li>
		<li>Capsule filters
		<ol>
			<li>Clean the capsule filters by first flushing them with de-ionized water, then 10% (v/v) HCl solution for a 48-hr soaking step.</li>
			<li>After acid soaking, flush the filters with de-ionized water and add 1 ml of 21% (weight) NH4OH (sub-boiled) solution into each filter to neutralize the acid.</li>
			<li>Seal individual filter with a loop of 30-cm cleaned-pumping tube connecting the inlet and outlet, and store in polyethylene zippered bags.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Sample containers
	<ol>
		<li>Use polyethylene (PE, 1,000-ml) and FEP (500 ml and 1,000 ml) bottles for containers for trace metal determination.
		<ol>
			<li>Clean the bottles by soaking them first in detergent (1%), then in 50% (v/v) HNO3 (Reagent Grade) and 10% (v/v) HCl (Reagent Grade) solutions for 24, 48 and 24 hr, respectively, and rinse the bottles with de-ionized water between the two soaking steps.</li>
			<li>After a final HCl soaking, rinse the bottles thoroughly with de-ionized water (DIW) and dry the bottles (with cap sealed) in a clean room or class-100 clean bench.</li>
			<li>Seal cleaned bottles in polyethylene zippered bags and double-bag them in polyethylene bags for transport.</li>
		</ol>
		</li>
		<li>Clean glass bottles for dissolved organic carbon (DOC) determination
		<ol>
			<li>Soak amber borosilicate glass bottles (40-ml), for analysis of dissolved organic carbon (DOC), in 10% HCl for 48 hr. Rinse cleaned glass bottles with de-ionized water, and combust them at 480 &#176;C for 2 hr before use. Seal the bottles individually in zippered polyethylene bags for transport.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Sampling
<ol>
	<li>Water sample collection
	<ol>
		<li>Upon arrival at the sampling site, mark sample identification number on the outer bag, and keep the bottles in their original bags.</li>
		<li>At the river bank or on a boat, have one person open the bag with the sampler and attach the 4-m FEP tubing to a 3-m polypropylene pole (cleaned). Extend the pole as far from the bank as possible and keep the inlet of the FEP tubing approximately 30 cm below the surface of running river water before the pump is turned on.</li>
		<li>Have another person attach the pumping tube to the pump head of a peristaltic pump (sampling pump with internal batteries). Start the pump and drain the water (downstream side) at least 3 times of the total sampler volume. Put on powder-free gloves and open the bag of bottles and caps to start filling the sample bottles.
		<ol>
			<li>Alternatively, if another person is available, have the third person be in charge of opening the inner bags and sample caps, and holding the sampling tube that drains the sample into the bottle.</li>
		</ol>
		</li>
		<li>Collect an unfiltered sample in a 125-ml plastic bottle for measurements of conductivity, temperature and pH in the field.</li>
		<li>Collect unfiltered samples (500-ml or 1,000 ml for particulate sample collection) first (through the outlet without a filter) and then close the outlet with a plastic clamp to force water to go through the capsule filter to collect a filtered water sample (1-L for dissolved trace metal determination).</li>
		<li>Collect filtered samples (through the outlet with the filter) in 40-ml amber glass bottles for DOC and EDTA measurements.</li>
		<li>Collect water samples at a flow rate of approximate 500 ml/min to 1,000 ml/min. Replace capsule filters when the pressure starts to build up (according to the filters&#39; specification). For each type of sample, collect extra samples, as well as field blanks, at selected locations to serve as quality control aliquots.</li>
		<li>Place the 40-ml glass bottles on dry ice and stored in an ice chest, and polyethylene bottles in ice chests.</li>
	</ol>
	</li>
	<li>Collection of suspended particulate matter (SPM)
	<ol>
		<li>Collect SPM on 0.4 &#956;m polycarbonate (PC) membrane filters (acid washed and pre-weighed) by vacuum filtration, using a plastic filtration funnel and vessel.</li>
		<li>Freeze dry membrane filters in the lab to yield SPM concentrations and provide particle samples for particulate trace metal determination.</li>
	</ol>
	</li>
</ol>
3. Sample Pretreatment (Dissolved Trace Metals)
<ol>
	<li>For dissolved trace metal determination, add 2-ml concentrated sub-boiled HNO3 (per 1-L sample) into sample bottles. Transfer dissolved trace metal samples (acidified) into FEP bottles. Alternately, samples can be collected directly into FEP bottles. UV-irradiate the samples in FEP bottles for 24 hr (8 15-watt UV lamps).</li>
</ol>
4. Preconcentration and Treatments for Trace Metal Analysis
<ol>
	<li>Preconcentration resin
	<ol>
		<li>Weigh 2 g of cation exchange resin into a small (10-30 ml capacity) plastic cup and add a small amount of 2 N HNO3 solution into the cup. Pour the resin into a 10-ml capacity chromatography column. Clean the resin by washing the column with 5-ml 2 N HNO3 (sub-boiled) twice, and rinse with high purity water (HPW) 3 times.</li>
		<li>Add 10 ml of 1 M NH4OH (sub-boiled) into the column to convert the resin to NH4+-form.</li>
	</ol>
	</li>
	<li>Buffer solution (1 M ammonium acetate)
	<ol>
		<li>Add 57 ml of glacial acetic acid (sub-boiled) in approximately 800 ml of HPW. Add ~60 ml of ammonium hydroxide (21%, sub-boiled) and mix with acetic acid. Adjust the final pH to 5.5 and the final volume to 1,000 ml.</li>
	</ol>
	</li>
	<li>Preconcentration procedure (dissolved trace metals)
	<ol>
		<li>Adjust the pH of the acidified, UV-irradiated samples to 5.5 &#177; 0.3 by adding 30 ml of 1 M ammonium acetate buffer solution and some (~2.8 ml) NH4OH (sub-boiled) into the samples.</li>
		<li>Place the sample bottle ~30 cm above the column packed with cationic exchange resin (Section 4.1) and connect the sample bottle and preconcentration column by a ~60 cm FEP tube and chromatography cap and connector (female Luer).</li>
		<li>Control the flow rate at 3-5 ml/min using a 2-way stopcock, connected above the column. Allow the samples to pass through the preconcentration columns. After the samples have passed the column, disconnect the tubes and caps from the columns.</li>
		<li>Treat the columns with 2 x 5-ml HPW and 4 x 5-ml of 1 M ammonium acetate (pH 5.5), and 2 x 5-ml of HPW to separate major cations from other trace metals.</li>
		<li>Place a 30-ml acid-washed PE bottle just below the column and wash the column with 7 x 1-ml 2 N HNO3 (sub-boiled) into the PE bottle.</li>
		<li>Determine the volume of 2 N HNO3 effluent (~8.0 ml) by obtaining the weight and specific gravity of the 2 N HNO3 effluent.</li>
	</ol>
	</li>
	<li>Digestion of suspended particulate matter
	<ol>
		<li>Freeze dry PC filters with SPM samples and weigh them after drying. Place SPM samples, with filters, into pre-weighed perfluoroalkoxy alkanes (PFA) vessels (60-ml capacity), and add 3 ml of concentrated HNO3 into the vessels.</li>
		<li>Tighten the vessels to a constant torque of 2.5 kg-meters, and digested in a conventional oven at 130 &#176;C for 12 hr. After cooling, open the vessels and add 2 ml of HF into the vessels.</li>
		<li>Tighten the vessels to a constant torque of 2.5 kg-meters, and digested in a conventional oven at 130 &#176;C for 12 hr. After cooling, open the vessels and add 16 ml of 4.5% boric acid solution into the vessels.</li>
		<li>Tighten the vessels to a constant torque of 2.5 kg-meters, and digest in a conventional oven at 130 &#176;C for 12 hr. Weigh each vessel and determine specific gravity of each digest solution to yield final digest volume.</li>
		<li>Calculate a dilution factor for each digest from the final volume and weight of SPM on filter (final digest volume divided by sample weight on filter).</li>
	</ol>
	</li>
</ol>
5. Sample Analysis
<ol>
	<li>Trace metals
	<ol>
		<li>Determine trace metals (Cd, Co, Cu, Fe, Mn, Ni, Pb, and Zn) concentrations in pre-concentrated dissolved sample and solution from particulates using flame atomic absorption spectrometry, graphite furnace atomic absorption spectrometry, and/or inductively coupled plasma mass spectrometer.</li>
		<li>Determine major ions concentrations and some trace metals, in sub-samples drawn prior to preconcentration and digests of suspended particulate matter, by an inductively coupled plasma-atomic emission spectrometry.</li>
	</ol>
	</li>
	<li>Ancillary parameters
	<ol>
		<li>Determine sample temperature, pH, salinity and conductivity in the field, using portable devices.</li>
		<li>Determine dissolved organic carbon (DOC) concentrations by a Total Organic Carbon Analyzer, based on wet oxidation with carbon dioxide detection by infrared spectrometry 21. Determine total EDTA concentration by a high performance liquid chromatography (with a SPD-M10AV Diodearray detector) following the established procedures 22,23.</li>
	</ol>
	</li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Preconcentration Techniques for Trace Elements. , 181-210 (1992).</li><li>Pai, S. -. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pre-concentration+efficiency+of+Chelex-100+resin+for+heavy+metals+in+seawater.+Part+2.+Distribution+of+heavy+metals+on+a+Chelex-100+column+and+optimization+of+the+column+efficiency+by+a+plate+simulation+method.">Pre-concentration efficiency of Chelex-100 resin for heavy metals in seawater. Part 2. Distribution of heavy metals on a Chelex-100 column and optimization of the column efficiency by a plate simulation method.</a> Analytica Chimica Acta. 211, 271-280 (1988).</li><li>Pai, S. -. C., Fang, T. -. H., Chen, C. -. T. A., Jeng, K. -. 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H., Salbu, B., Steinnes, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Trace Elements in Natural Waters. , 41-69 (1995).</li><li>. U.S. Environmental Protection Agency. Method 1669. Sampling ambient water for trace metals at EPA Water Quality criteria levels Available from: <a target="_blank" href="https://www3.epa.gov/caddis/pdf/Metals_Sampling_EPA_method_1669.pdf">https://www3.epa.gov/caddis/pdf/Metals_Sampling_EPA_method_1669.pdf</a> (1996)</li><li>Horowitz, A. 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P., Armstrong, D., Webb, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+influence+of+dissolved+organic+carbon,+suspended+particles,+and+hydrology+on+the+concentration,+partitioning+and+variability+of+trace+metals+in+two+contrasting+Wisconsin+watersheds+(U.S.A.).">The influence of dissolved organic carbon, suspended particles, and hydrology on the concentration, partitioning and variability of trace metals in two contrasting Wisconsin watersheds (U.S.A.).</a> Chemical Geology. 136, 71-97 (1997).</li><li>Warren, L. A., Haack, E. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biogeochemical+controls+on+metal+behaviour+in+freshwater+environments.">Biogeochemical controls on metal behaviour in freshwater environments.</a> Earth-Science Reviews. 54, 261-320 (2001).</li><li>Jiann, K. -. T., Santschi, P. H., Presley, B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Relationships+between+geochemical+parameters+(pH,+DOC,+SPM,+EDTA+Concentrations)+and+trace+metal+(Cd,+Co,+Cu,+Fe,+Mn,+Ni,+Pb,+Zn)+concentrations+in+river+waters+of+Texas+(USA).">Relationships between geochemical parameters (pH, DOC, SPM, EDTA Concentrations) and trace metal (Cd, Co, Cu, Fe, Mn, Ni, Pb, Zn) concentrations in river waters of Texas (USA).</a> Aquatic Geochemistry. 19 (2), 173-193 (2013).</li><li>Peltzer, E. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparison+of+methods+for+the+measurement+of+dissolved+organic+carbon+in+natural+waters.">A comparison of methods for the measurement of dissolved organic carbon in natural waters.</a> Marine Chemistry. 54, 85-96 (1996).</li><li>Nowack, B., Kari, F., Hilger, S. U., Sigg, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determination+of+dissolved+and+adsorbed+EDTA+species+in+water+and+sediments+by+HPLC.">Determination of dissolved and adsorbed EDTA species in water and sediments by HPLC.</a> Analytical Chemistry. 68 (3), 561-566 (1996).</li><li>Bergers, P. J. M., de Groot, A. 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The authors have no conflict of interests to disclose.

Obtaining reliable trace metal data in natural waters requires great care as emphasized during sample collection, processing, pretreatments, and analysis that aim to reduce contamination. Trace metal concentrations in natural waters obtained using &#34;clean techniques&#34; in the last two decades found that the concentrations can be orders of magnitude lower than previously reported. Water quality criteria for trace metals in waters are now more readily assessed when trace metal levels are accurately measured resulting ...

It has been commonly recognized that some trace metal results obtained for natural waters may be inaccurate owing to artifacts arising from inadequate techniques applied during sample collection, treatments and determination1,2. The true concentrations (in sub-nM to nM range in surface waters 3) of dissolved trace metals are now up to two orders of magnitude lower than previously published values. The same situation has been found in marine chemistry where the accepted dissolved trace metal concentrations in oceanic waters have decreased by orders of magnitude over the last 40 years or so as improved sampling and analytical methods have been introduced. Efforts have been made to improve data quality with the developments of &#34;clean techniques&#34; aiming at the reduction or elimination of trace metal contamination throughout all phases of trace metal analysis 4-8. For the determination of trace metal concentrations at ambient levels, preconcentration is often required. Ion-exchange techniques 8-12 have been commonly applied for efficient preconcentration.
Contamination can arise from the walls of containers, the cleaning of the containers, the sampler, sample handling and storage, and sample preservation and analysis 7,13. All studies using clean methods conducted more recently indicate that trace metal concentrations in natural waters are typically well below the detection limits of routine methods 7. Since the recognition of suspect trace metal data in the early 1990s, clean methods have been incorporated into US EPA (Environmental Protection Agency) Guidelines for trace metal determination 14 and the US Geological Survey has adopted clean methods for their water quality monitoring projects 15. Clean methods for trace metal studies need to be employed in all projects in order to create a firm and accurate data base.
In principle, water samples used for trace metal determination should be collected with appropriate sampling gears of a particular material and composition, stored and treated properly using appropriate containers and apparatus, before proceeding with instrumental analysis. Since suspended particulate matter (SPM) can undergo changes during the sample storage period and alter water composition, rapid separation of SPM from water samples is a common practice for trace metal studies in aquatic environments. For the determination of dissolved trace metal concentrations in natural waters, filtration is necessary and in-line filtration techniques are suitable and efficient.
Distribution and behavior of trace metals in aquatic environments such as surface and ground waters can be affected by natural (e.g., weathering) and anthropogenic (e.g., wastewater effluents) factors, as well as other environmental conditions, such as regional geology, morphology, land use and vegetation, and climate 16-19. This can then lead to differences in physicochemical parameters such as concentrations of suspended particulate matter (SPM), dissolved organic carbon (DOC), anthropogenic ligands (e.g., ethylenediaminetetraacetic acid, EDTA), salt, redox potential and pH 17-20. Therefore, accurate and relevant trace metal studies require appropriate collection of samples for trace metal analysis as well as for the determination of related factors and parameters.

<table><tbody><tr><td>Nitric Acid</td><td>Seastar Chemicals</td><td></td><td>Baseline grade</td></tr><tr><td>Ammonium hydroxide</td><td>Seastar Chemicals</td><td></td><td>Baseline grade</td></tr><tr><td>Acetic Acid</td><td>Seastar Chemicals</td><td></td><td>Baseline grade</td></tr><tr><td>Nitric Acid</td><td>J. T. Baker</td><td>9601-05</td><td>Reagent grade</td></tr><tr><td>Hydrochloric acid</td><td>J. T. Baker</td><td>9530-33</td><td>Reagent grade</td></tr><tr><td>Chromatographic columns</td><td>Bio-Rad</td><td>7311550&amp;nbsp;</td><td>Poly-Prep</td></tr><tr><td>Column stack caps</td><td>Bio-Rad</td><td>7311555</td><td></td></tr><tr><td>Cap connectors (female Luers)</td><td>Bio-Rad</td><td>7318223</td><td></td></tr><tr><td>2-way stopcocks</td><td>Bio-Rad</td><td>7328102</td><td></td></tr><tr><td>Cation exchange resin</td><td>Bio-Rad</td><td>1422832&amp;nbsp;</td><td>Chelex-100</td></tr><tr><td>Portable sampler (sampling pump)</td><td>Cole Palmer</td><td>EW-07571-00</td><td></td></tr><tr><td>FEP tube</td><td>Cole Palmer</td><td>EW-06450-07</td><td>6.4 mm I.D., 9.5 mm O.D.</td></tr><tr><td>Pumping tube</td><td>Cole Palmer</td><td>EW-06424-24</td><td>6.4 mm I.D. C-Flex</td></tr><tr><td>Capsule filter (0.4 mm)</td><td>Fisher Scientific</td><td>WP4HY410F0</td><td>polypropylene casing</td></tr><tr><td>1 L low density polyethylene bottle</td><td>NALGE NUNC INTERNATIONAL</td><td>312088-0032</td><td></td></tr><tr><td>1 L (or 500 ml) FEP bottle</td><td>NALGE NUNC INTERNATIONAL</td><td>381600-0032</td><td></td></tr></tbody></table>

clean sampling and analysis of river and estuarine waters for trace metal studies

Most of the trace metal concentrations in ambient waters obtained a few decades ago have been considered unreliable owing to the lack of contamination control. Developments of some techniques aiming to reduce trace metal contamination in the last couple of decades have resulted in concentrations reported now being orders of magnitude lower than those in the past. These low concentrations often necessitate preconcentration of water samples prior to instrumental analysis of samples. Since contamination can appear in all phases of trace metal analyses, including sample collection (and during preparation of sampling containers), storage and handling, pretreatments, and instrumental analysis, specific care needs to be taken in order to reduce contamination levels at all steps. The effort to develop and utilize "clean techniques" in trace metal studies allows scientists to investigate trace metal distributions and chemical and biological behavior in greater details. This advancement also provides the required accuracy and precision of trace metal data allowing for environmental conditions to be related to trace metal concentrations in aquatic environments.
This protocol that is presented here details needed materials for sample preparation, sample collection, sample pretreatment including preconcentration, and instrumental analysis. By reducing contamination throughout all phases mentioned above for trace metal analysis, much lower detection limits and thus accuracy can be achieved. The effectiveness of "clean techniques" is further demonstrated using low field blanks and good recoveries for standard reference material. The data quality that can be obtained thus enables the assessment of trace metal distributions and their relationships to environmental parameters.

With the development and use of &#34;clean techniques&#34;, it is now well recognized that in order to obtain accurate trace metal concentrations in ambient waters, preconcentration of trace metals in waters samples is a common practice. While most water quality criteria for trace metals in natural waters are in the low &#956;g/L range, lower detection limits are needed to investigate geochemical and biological effects on trace metals at ambient concentrations in aquatic environments....

Watch this Scientific Journal Video about Clean Sampling and Analysis of River and Estuarine Waters for Trace Metal Studies at JoVE.com

Clean Sampling and Analysis of River and Estuarine Waters for Trace Metal Studies

Special care using "clean techniques" is required to properly collect and process water samples for trace metal studies in aquatic environments. A protocol for sampling, processing, and analytical procedures with the aim of obtaining reliable environmental monitoring data and results with high sensitivity for detailed trace metal studies is presented.

Most of the trace metal concentrations in ambient waters obtained a few decades ago have been considered unreliable owing to the lack of contamination ...

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Research

JoVE Journal

Environment

11.3K Views.  National Sun Yat-sen University. Special care using "clean techniques" is required to properly collect and process water samples for trace metal studies in aquatic environments. A protocol for sampling, processing, and analytical procedures with the aim of obtaining reliable environmental monitoring data and results with high sensitivity for detailed trace metal studies is presented.

Video: Clean Sampling and Analysis of River and Estuarine Waters for Trace Metal Studies

Quantification of Heavy Metals and Other Inorganic Contaminants on the Productivity of Microalgae

Increasing demand for renewable fuels has researchers investigating the feasibility of alternative feedstocks, such as microalgae. Inherent advantages include high potential yield, use of non-arable land and integration with waste streams. The nutrient requirements of a large-scale microalgae production system will require the coupling of cultivation systems with industrial waste resources, such as carbon dioxide from flue gas and nutrients from wastewater. Inorganic contaminants present in these wastes can potentially lead to bioaccumulation in microalgal biomass negatively impact productivity and limiting end use. This study focuses on the experimental evaluation of the impact and the fate of 14 inorganic contaminants (As, Cd, Co, Cr, Cu, Hg, Mn, Ni, Pb, Sb, Se, Sn, V and Zn) on Nannochloropsis salina growth. Microalgae were cultivated in photobioreactors illuminated at 984 &#181;mol m-2 sec-1 and maintained at pH 7 in a growth media polluted with inorganic contaminants at levels expected based on the composition found in commercial coal flue gas systems. Contaminants present in the biomass and the medium at the end of a 7 day growth period were analytically quantified through cold vapor atomic absorption spectrometry for Hg and through inductively coupled plasma mass spectrometry for As, Cd, Co, Cr, Cu, Mn, Ni, Pb, Sb, Se, Sn, V and Zn. Results show N. salina is a sensitive strain to the multi-metal environment with a statistical decrease in biomass yieldwith the introduction of these contaminants. The techniques presented here are adequate for quantifying algal growth and determining the fate of inorganic contaminants.

Integration of microalgal cultivation with industrial flue gas will ultimately introduce heavy metals and other inorganic compounds into the growth media. This study presents a procedure used to determine the end fate and impact of heavy metals and inorganic contaminants on the growth of Nannochloropsis salina grown in photobioreactors.

Increasing demand for renewable fuels has researchers investigating the feasibility of alternative feedstocks, such as microalgae. Inherent advantages ...

Sediment Core Sectioning and Extraction of Pore Waters under Anoxic Conditions

We demonstrate a method for sectioning sediment cores and extracting pore waters while maintaining oxygen-free conditions. A simple, inexpensive system is built and can be transported to a temporary work space close to field sampling site(s) to facilitate rapid analysis. Cores are extruded into a portable glove bag, where they are sectioned and each 1-3 cm thick section (depending on core diameter) is sealed into 50 ml centrifuge tubes. Pore waters are separated with centrifugation outside of the glove bag and then returned to the glove bag for separation from the sediment. These extracted pore water samples can be analyzed immediately. Immediate analyses of redox sensitive species, such as sulfide, iron speciation, and arsenic speciation indicate that oxidation of pore waters is minimal; some samples show approximately 100% of the reduced species, e.g. 100% Fe(II) with no detectable Fe(III). Both sediment and pore water samples can be preserved to maintain chemical species for further analysis upon return to the laboratory.

A protocol for sectioning sediment cores and extracting pore waters under anoxic conditions in order to permit analysis of redox sensitive species in both solids and fluids is presented.

We demonstrate a method for sectioning sediment cores and extracting pore waters while maintaining oxygen-free conditions. A simple, inexpensive system is ...

Watershed Planning within a Quantitative Scenario Analysis Framework

There is a critical need for tools and methodologies capable of managing aquatic systems within heavily impacted watersheds. Current efforts often fall short as a result of an inability to quantify and predict complex cumulative effects of current and future land use scenarios at relevant spatial scales. The goal of this manuscript is to provide methods for conducting a targeted watershed assessment that enables resource managers to produce landscape-based cumulative effects models for use within a scenario analysis management framework. Sites are first selected for inclusion within the watershed assessment by identifying sites that fall along independent gradients and combinations of known stressors. Field and laboratory techniques are then used to obtain data on the physical, chemical, and biological effects of multiple land use activities. Multiple linear regression analysis is then used to produce landscape-based cumulative effects models for predicting aquatic conditions. Lastly, methods for incorporating cumulative effects models within a scenario analysis framework for guiding management and regulatory decisions (e.g., permitting and mitigation) within actively developing watersheds are discussed and demonstrated for 2 sub-watersheds within the mountaintop mining region of central Appalachia. The watershed assessment and management approach provided herein enables resource managers to facilitate economic and development activity while protecting aquatic resources and producing opportunity for net ecological benefits through targeted remediation.

There is a critical need for tools and methodologies capable of managing aquatic systems in the face of uncertain future conditions. We provide methods for conducting a targeted watershed assessment that enables resource managers to produce landscape-based cumulative effects models for use within a scenario analysis management framework.

There is a critical need for tools and methodologies capable of managing aquatic systems within heavily impacted watersheds. Current efforts often fall ...

Continuous Instream Monitoring of Nutrients and Sediment in Agricultural Watersheds

Pollutant concentrations and loads in watersheds vary considerably with time and space. Accurate and timely information on the magnitude of pollutants in water resources is a prerequisite for understanding the drivers of the pollutant loads and for making informed water resource management decisions. The commonly used &#34;grab sampling&#34; method provides the concentrations of pollutants at the time of sampling (i.e., a snapshot concentration) and may under- or overpredict the pollutant concentrations and loads. Continuous monitoring of nutrients and sediment has recently received more attention due to advances in computing, sensing technology, and storage devices. This protocol demonstrates the use of sensors, sondes, and instrumentation to continuously monitor in situ nitrate, ammonium, turbidity, pH, conductivity, temperature, and dissolved oxygen (DO) and to calculate the loads from two streams (ditches) in two agricultural watersheds. With the proper calibration, maintenance, and operation of sensors and sondes, good water quality data can be obtained by overcoming challenging conditions such as fouling and debris buildup. The method can also be used in watersheds of various sizes and characterized by agricultural, forested, and/or urban land.

With the advancement of technology and the rise in end-user expectations, the need and use of higher temporal resolution data for pollutant load estimation has increased. This protocol describes a method for continuous in situ water quality monitoring to obtain higher temporal resolution data for informed water resource management decisions.

Pollutant concentrations and loads in watersheds vary considerably with time and space. Accurate and timely information on the magnitude of pollutants in ...

An Ultra-clean Multilayer Apparatus for Collecting Size Fractionated Marine Plankton and Suspended Particles

The distributions of many trace elements in the ocean are strongly associated with the growth, death, and re-mineralization of marine plankton and those of suspended/sinking particles. Here, we present an all plastic (Polypropylene and Polycarbonate), multi-layer filtration system for collection of suspended particulate matter (SPM) at sea. This ultra-clean sampling device has been designed and developed specifically for trace element studies. Meticulous selection of all non-metallic materials and utilization of an in-line flow-through procedure minimizes any possible metal contamination during sampling. This system has been successfully tested and tweaked for determining trace metals (e.g., Fe, Al, Mn, Cd, Cu, Ni) on particles of varying size in coastal and open ocean waters. Results from the South China Sea at the South East Asia Time-Series (SEATS) station indicate that diurnal variations and spatial distribution of plankton in the euphotic zone can be easily resolved and recognized. Chemical analysis of size-fractionated particles in surface waters of the Taiwan Strait suggests that the larger particles (&#62;153 &#181;m) were mostly biologically derived, while the smaller particles (10 - 63 &#181;m) were mostly composed of inorganic matter. Apart from Cd, the concentrations of metals (Fe, Al, Mn, Cu, Ni) decreased with increasing size.

Plankton and suspended particles play a major role in the biogeochemical cycles in the ocean. Here, we provide an ultra-clean, low stress method for the collection of various sizes of particles and plankton at sea with the capability of handling large volumes of seawater.

The distributions of many trace elements in the ocean are strongly associated with the growth, death, and re-mineralization of marine plankton and those ...

Seawater Sampling and Collection

This video documents methods for collecting coastal marine water samples and processing them for various downstream applications including biomass concentration, nucleic acid purification, cell abundance, nutrient and trace gas analyses. For today's demonstration samples were collected from the deck of the HMS John Strickland operating in Saanich Inlet. An A-frame derrick, with a multi-purpose winch and cable system, is used in combination with Niskin or Go-Flo water sampling bottles. Conductivity, Temperature, and Depth (CTD) sensors are also used to sample the underlying water mass. To minimize outgassing, trace gas samples are collected first.  Then, nutrients, water chemistry, and cell counts are determined. Finally, waters are collected for biomass filtration. The set-up and collection time for a single cast is ~1.5 hours at a maximum depth of 215 meters. Therefore, a total of 6 hours is generally needed to complete the collection series described here.

This video documents methods for collecting coastal marine water samples and processing them for various downstream applications including biomass concentration, nucleic acid purification, cell abundance, nutrient and trace gas analyses.

This video documents methods for collecting coastal marine water samples and processing them for various downstream applications including biomass ...

Biology

Two-Dimensional Visualization and Quantification of Labile, Inorganic Plant Nutrients and Contaminants in Soil

We describe a method for two-dimensional (2D) visualization and quantification of the distribution of labile (i.e., reversibly adsorbed) inorganic nutrient (e.g., P, Fe, Mn) and contaminant (e.g., As, Cd, Pb) solute species in the soil adjacent to plant roots (the &#8216;rhizosphere&#8217;) at sub-millimeter (~100 &#181;m) spatial resolution. The method combines sink-based solute sampling by the diffusive gradients in thin films (DGT) technique with spatially resolved chemical analysis by laser ablation inductively coupled plasma mass spectrometry (LA-ICP-MS). The DGT technique is based on thin hydrogels with homogeneously distributed analyte-selective binding phases. The variety of available binding phases allows for the preparation of different DGT gel types following simple gel fabrication procedures. For DGT gel deployment in the rhizosphere, plants are grown in flat, transparent growth containers (rhizotrons), which enable minimal invasive access to a soil-grown root system. After a pre-growth period, DGT gels are applied to selected regions of interest for in situ solute sampling in the rhizosphere. Afterwards, DGT gels are retrieved and prepared for subsequent chemical analysis of the bound solutes using LA-ICP-MS line-scan imaging. Application of internal normalization using 13C and external calibration using matrix-matched gel standards further allows for the quantification of the 2D solute fluxes. This method is unique in its capability to generate quantitative, sub-mm scale 2D images of multi-element solute fluxes in soil-plant environments, exceeding the achievable spatial resolution of other methods for measuring solute gradients in the rhizosphere substantially. We present the application and evaluation of the method for imaging multiple cationic and anionic solute species in the rhizosphere of terrestrial plants and highlight the possibility of combining this method with complementary solute imaging techniques.

This protocol presents a workflow for sub-mm 2D visualization of multiple labile inorganic nutrient and contaminant solute species using diffusive gradients in thin films (DGT) combined with mass spectrometry imaging. Solute sampling and high-resolution chemical analysis are described in detail for quantitative mapping of solutes in the rhizosphere of terrestrial plants.

We describe a method for two-dimensional (2D) visualization and quantification of the distribution of labile (i.e., reversibly adsorbed) inorganic ...

Dissolved Solute Sampling Across an Oxic-Anoxic Soil-Water Interface Using Microdialysis Profilers

Biogeochemical processes shift rapidly in both spatial (millimeter scale) and temporal (hour scale to day scale) dimensions at the oxic-anoxic interface in response to disturbances. Deciphering the rapid biogeochemical changes requires in situ, minimally invasive tools with high spatial and temporal sampling resolution. However, the available passive sampling devices are not very useful in many cases either due to their disposable nature or the complexity and extensive workload for sample preparation.
To address this problem, a microdialysis profiler with 33 individual polyethersulfone nanomembrane tubes (semipermeable, &#60;20 nm pore size) integrated into the one-dimensional skeleton (60 mm) was established to iteratively sample the dissolved compounds in porewater across the soil-water interface at a high resolution of 1.8 mm (outer diameter plus one spacing, i.e., 0.1 mm between probes). The sampling mechanism is based on the principle of concentration gradient diffusion. The automatic loading of degassed water allows minimal disturbance to the chemical species across the oxic-anoxic interface.
This paper describes the procedures of device setup and continuous porewater sampling across the soil-water interface on a daily basis. Concentration-depth profiles were selectively measured before (on Day 6) and after (on Day 7) disturbances induced by irrigation. The results showed that concentration-depth profiles were undergoing rapid changes, especially for redox-sensitive elements (i.e., iron and arsenic). These protocols can help investigate the biogeochemical responses across the soil-water interface under various disturbances caused by physical, chemical, and biological factors. The paper thoroughly discusses the advantages and disadvantages of this method for potential use in the environmental sciences.

A microdialysis profiler is described to sample dissolved porewater solutes across an oxic-anoxic soil-water interface in situ with minimal disturbance. This device is designed to capture rapid changes in concentration-depth profiles induced by disturbances at the soil-water interface and beyond.

Biogeochemical processes shift rapidly in both spatial (millimeter scale) and temporal (hour scale to day scale) dimensions at the oxic-anoxic interface ...

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 ...

Fabrication of a Dipole-assisted Solid Phase Extraction Microchip for Trace Metal Analysis in Water Samples

This paper describes a fabrication protocol for a dipole-assisted solid phase extraction (SPE) microchip available for trace metal analysis in water samples. A brief overview of the evolution of chip-based SPE techniques is provided. This is followed by an introduction to specific polymeric materials and their role in SPE. To develop an innovative dipole-assisted SPE technique, a chlorine (Cl)-containing SPE functionality was implanted into a poly(methyl methacrylate) (PMMA) microchip. Herein, diverse analytical techniques including contact angle analysis, Raman spectroscopic analysis, and laser ablation-inductively coupled plasma-mass spectrometry (LA-ICP-MS) analysis were employed to validate the utility of the implantation protocol of the C-Cl moieties on the PMMA. The analytical results of the X-ray absorption near-edge structure (XANES) analysis also demonstrated the feasibility of the Cl-containing PMMA used as an extraction medium by virtue of the dipole-ion interactions between the highly electronegative C-Cl moieties and the positively charged metal ions.

The fabrication protocol of a dipole-assisted solid phase extraction microchip for the trace metal analysis is presented.

This paper describes a fabrication protocol for a dipole-assisted solid phase extraction (SPE) microchip available for trace metal analysis in water ...