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==Sediment Capping==
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==Sediment Porewater Dialysis Passive Samplers for Inorganics (Peepers)==  
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Capping® is an ''in situ'' remedial technology that involves placement of a clean substrate on the surface of [[Contaminated Sediments - Introduction | contaminated sediments]] to reduce contaminant uptake by benthic organisms and contaminant flux to surface water. Simple sand caps can be effective in reducing exposure of benthic organisms and in limiting oxygen transport into the contaminated sediments, resulting in precipitation of metal sulfides. Amendments are sometimes included in caps to reduce cap permeability and groundwater upwelling, to increase contaminant sorption or biodegradation, or to improve habitat.
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Sediment porewater dialysis passive samplers, also known as “peepers,” are sampling devices that allow the measurement of dissolved inorganic ions in the porewater of a saturated sediment. Peepers function by allowing freely-dissolved ions in sediment porewater to diffuse across a micro-porous membrane towards water contained in an isolated compartment that has been inserted into sediment. Once retrieved after a deployment period, the resulting sample obtained can provide concentrations of freely-dissolved inorganic constituents in sediment, which provides measurements that can be used for understanding contaminant fate and risk. Peepers can also be used in the same manner in surface water, although this article is focused on the use of peepers in sediment.  
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<div style="float:right;margin:0 0 2em 2em;">__TOC__</div>
 
<div style="float:right;margin:0 0 2em 2em;">__TOC__</div>
  
 
'''Related Article(s):'''
 
'''Related Article(s):'''
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*[[Contaminated Sediments - Introduction]]
 
*[[Contaminated Sediments - Introduction]]
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*[[Contaminated Sediment Risk Assessment]]
 
*[[In Situ Treatment of Contaminated Sediments with Activated Carbon]]
 
*[[In Situ Treatment of Contaminated Sediments with Activated Carbon]]
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*Sediment Risk Assessment
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*[[Passive Sampling of Munitions Constituents]]
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*[[Sediment Capping]]
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*[[Mercury in Sediments]]
 
*[[Passive Sampling of Sediments]]
 
*[[Passive Sampling of Sediments]]
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'''Contributor(s):'''  
 
'''Contributor(s):'''  
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*Danny Reible
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*Florent Risacher, M.Sc.
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*Jason Conder, Ph.D.
  
 
'''Key Resource(s):'''
 
'''Key Resource(s):'''
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*Processes, Assessment and Remediation of Contaminated Sediments<ref name="Reible2014">Reible, D. D., Editor, 2014. Processes, Assessment and Remediation of Contaminated Sediments. Springer, New York, NY. 462 pp. ISBN: 978-1-4614-6725-0</ref>
 
  
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* Guidance for In-Situ Subaqueous Capping of Contaminated Sediments<ref name="Palermo1998">Palermo, M., Maynord, S., Miller, J. and Reible, D., 1998. Guidance for In-Situ Subaqueous Capping of Contaminated Sediments. Assessment and Remediation of Contaminated Sediments (ARCS) Program, Great Lakes National Program Office, US EPA 905-B96-004. 147 pp.  [[Media: USEPA_905-B96-004.pdf | Report.pdf]]</ref>
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*A review of peeper passive sampling approaches to measure the availability of inorganics in sediment porewater<ref>Risacher, F.F., Schneider, H., Drygiannaki, I., Conder, J., Pautler, B.G., and Jackson, A.W., 2023.  A Review of Peeper Passive Sampling Approaches to Measure the Availability of Inorganics in Sediment Porewater.  Environmental Pollution, 328, Article 121581. [https://doi.org/10.1016/j.envpol.2023.121581 doi: 10.1016/j.envpol.2023.121581]&nbsp;&nbsp;[[Media: RisacherEtAl2023a.pdf | Open Access Manuscript]]</ref>
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*Best Practices User’s Guide: Standardizing Sediment Porewater Passive Samplers for Inorganic Constituents of Concern<ref name="RisacherEtAl2023">Risacher, F.F., Nichols, E., Schneider, H., Lawrence, M., Conder, J., Sweett, A., Pautler, B.G., Jackson, W.A., Rosen, G., 2023b. Best Practices User’s Guide: Standardizing Sediment Porewater Passive Samplers for Inorganic Constituents of Concern, ESTCP ER20-5261. [https://serdp-estcp.mil/projects/details/db871313-fbc0-4432-b536-40c64af3627f Project Website]&nbsp;&nbsp;[[Media: ER20-5261BPUG.pdf | Report.pdf]]</ref>
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*[https://serdp-estcp.mil/projects/details/db871313-fbc0-4432-b536-40c64af3627f/er20-5261-project-overview Standardizing Sediment Porewater Passive Samplers for Inorganic Constituents of Concern, ESTCP Project ER20-5261]
  
 
==Introduction==
 
==Introduction==
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[[File:SedCapFig1.png|thumb|left|470px|Figure 1. Conceptual sketch of a cap configuration]]
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Biologically available inorganic constituents associated with sediment toxicity can be quantified by measuring the freely-dissolved fraction of contaminants in the porewater<ref>Conder, J.M., Fuchsman, P.C., Grover, M.M., Magar, V.S., Henning, M.H., 2015. Critical review of mercury SQVs for the protection of benthic invertebrates. Environmental Toxicology and Chemistry, 34(1), pp. 6-21. [https://doi.org/10.1002/etc.2769 doi: 10.1002/etc.2769]&nbsp;&nbsp; [[Media: ConderEtAl2015.pdf | Open Access Article]]</ref><ref name="ClevelandEtAl2017">Cleveland, D., Brumbaugh, W.G., MacDonald, D.D., 2017. A comparison of four porewater sampling methods for metal mixtures and dissolved organic carbon and the implications for sediment toxicity evaluations. Environmental Toxicology and Chemistry, 36(11), pp. 2906-2915. [https://doi.org/10.1002/etc.3884 doi: 10.1002/etc.3884]</ref>. Classical sediment porewater analysis usually consists of collecting large volumes of bulk sediments which are then mechanically squeezed or centrifuged to produce a supernatant, or suction of porewater from intact sediment, followed by filtration and collection<ref name="GruzalskiEtAl2016">Gruzalski, J.G., Markwiese, J.T., Carriker, N.E., Rogers, W.J., Vitale, R.J., Thal, D.I., 2016. Pore Water Collection, Analysis and Evolution: The Need for Standardization. In: Reviews of Environmental Contamination and Toxicology, Vol. 237, pp. 37–51. Springer. [https://doi.org/10.1007/978-3-319-23573-8_2 doi: 10.1007/978-3-319-23573-8_2]</ref>. The extraction and measurement processes present challenges due to the heterogeneity of sediments, physical disturbance, high reactivity of some complexes, and interaction between the solid and dissolved phases, which can impact the measured concentration of dissolved inorganics<ref>Peijnenburg, W.J.G.M., Teasdale, P.R., Reible, D., Mondon, J., Bennett, W.W., Campbell, P.G.C., 2014. Passive Sampling Methods for Contaminated Sediments: State of the Science for Metals. Integrated Environmental Assessment and Management, 10(2), pp. 179–196. [https://doi.org/10.1002/ieam.1502 doi: 10.1002/ieam.1502]&nbsp;&nbsp; [[Media: PeijnenburgEtAl2014.pdf | Open Access Article]]</ref>. For example, sampling disturbance can affect redox conditions<ref name="TeasdaleEtAl1995">Teasdale, P.R., Batley, G.E., Apte, S.C., Webster, I.T., 1995. Pore water sampling with sediment peepers. Trends in Analytical Chemistry, 14(6), pp. 250–256. [https://doi.org/10.1016/0165-9936(95)91617-2 doi: 10.1016/0165-9936(95)91617-2]</ref><ref>Schroeder, H., Duester, L., Fabricius, A.L., Ecker, D., Breitung, V., Ternes, T.A., 2020. Sediment water (interface) mobility of metal(loid)s and nutrients under undisturbed conditions and during resuspension. Journal of Hazardous Materials, 394, Article 122543. [https://doi.org/10.1016/j.jhazmat.2020.122543 doi: 10.1016/j.jhazmat.2020.122543]&nbsp;&nbsp; [[Media: SchroederEtAl2020.pdf | Open Access Article]]</ref>, which can lead to under or over representation of inorganic chemical concentrations relative to the true dissolved phase concentration in the sediment porewater<ref>Wise, D.E., 2009. Sampling techniques for sediment pore water in evaluation of reactive capping efficacy. Master of Science Thesis. University of New Hampshire Scholars’ Repository. 178 pages. [https://scholars.unh.edu/thesis/502 Website]&nbsp;&nbsp; [[Media: Wise2009.pdf | Report.pdf]]</ref><ref name="GruzalskiEtAl2016"/>.
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Capping is an ''in situ'' remedial technology for contaminated sediments that involves placement of a clean substrate on the sediment surface.  Capping contaminated sediments following [[Wikipedia: Dredging | dredging operations]] and capping of dredged material to stabilize contaminants has been a common practice by the United States Army Corps of Engineers since the 1970s. Beginning in the 1980s, in Japan and subsequently elsewhere, capping has been used more widely as a remedial approach to improve the quality of the bottom substrate and reduce contaminant exposures to benthic organisms and fish. The USEPA published a capping guidance document in 1998 that summarizes past uses of sediment capping and outlines its basic design<ref name="Palermo1998"/>.   Although capping technology has developed substantially in the past 20 years, this early reference still provides useful information on the approach and its applications.  A more recent summary of capping is described in Reible 2014<ref name="Reible2014"/>.
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To address the complications with mechanical porewater sampling, passive sampling approaches for inorganics have been developed to provide a method that has a low impact on the surrounding geochemistry of sediments and sediment porewater, thus enabling more precise measurements of inorganics<ref name="ClevelandEtAl2017"/>. Sediment porewater dialysis passive samplers, also known as “peepers,” were developed more than 45 years ago<ref name="Hesslein1976">Hesslein, R.H., 1976. An in situ sampler for close interval pore water studies. Limnology and Oceanography, 21(6), pp. 912-914. [https://doi.org/10.4319/lo.1976.21.6.0912 doi: 10.4319/lo.1976.21.6.0912]&nbsp;&nbsp; [[Media: Hesslein1976.pdf | Open Access Article]]</ref> and refinements to the method such as the use of reverse tracers have been made, improving the acceptance of the technology as decision-making tool.
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==Peeper Designs==
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Peepers (Figure 1) are inert containers with a small volume (typically 1-100 mL) of purified water (“peeper water”) capped with a semi-permeable membrane. Peepers can be manufactured in a wide variety of formats (Figure 2, Figure 3) and deployed in in various ways.
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Two designs are commonly used for peepers. Frequently, the designs are close adaptations of the original multi-chamber Hesslein design<ref name="Hesslein1976"/> (Figure 2), which consists of an acrylic sampler body with multiple sample chambers machined into it. Peeper water inside the chambers is separated from the outside environment by a semi-permeable membrane, which is held in place by a top plate fixed to the sampler body using bolts or screws. An alternative design consists of single-chamber peepers constructed using a single sample vial with a membrane secured over the mouth of the vial, as shown in Figure 3, and applied in Teasdale ''et al.''<ref name="TeasdaleEtAl1995"/>, Serbst ''et al.''<ref>Serbst, J.R., Burgess, R.M., Kuhn, A., Edwards, P.A., Cantwell, M.G., Pelletier, M.C., Berry, W.J., 2003. Precision of dialysis (peeper) sampling of cadmium in marine sediment interstitial water. Archives of Environmental Contamination and Toxicology, 45(3), pp. 297–305. [https://doi.org/10.1007/s00244-003-0114-5 doi: 10.1007/s00244-003-0114-5]</ref>, Thomas and Arthur<ref name="ThomasArthur2010">Thomas, B., Arthur, M.A., 2010. Correcting porewater concentration measurements from peepers: Application of a reverse tracer. Limnology and Oceanography: Methods, 8(8), pp. 403–413. [https://doi.org/10.4319/lom.2010.8.403 doi: 10.4319/lom.2010.8.403]&nbsp;&nbsp; [[Media: ThomasArthur2010.pdf | Open Access Article]]</ref>, Passeport ''et al.''<ref>Passeport, E., Landis, R., Lacrampe-Couloume, G., Lutz, E.J., Erin Mack, E., West, K., Morgan, S., Lollar, B.S., 2016. Sediment Monitored Natural Recovery Evidenced by Compound Specific Isotope Analysis and High-Resolution Pore Water Sampling. Environmental Science and Technology, 50(22), pp. 12197–12204. [https://doi.org/10.1021/acs.est.6b02961 doi: 10.1021/acs.est.6b02961]</ref>, and Risacher ''et al.''<ref name="RisacherEtAl2023"/>. The vial is filled with deionized water, and the membrane is held in place using the vial cap or an o-ring. Individual vials are either directly inserted into sediment or are incorporated into a support structure to allow multiple single-chamber peepers to be deployed at once over a given depth profile (Figure 3).
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Capping serves to contain contaminated sediment solids, isolate contaminants from benthic organisms and reduce contaminant transport to the sediment surface and overlying water. The clean substrate may be an inert material such as sand, a natural sorbing material such as other sediments or clays, or be amended with an active/reactive material to enhance the isolation of the contaminants.  Amendments to enhance contaminant isolation include permeability reduction agents to divert groundwater flow, sorbents to retard contaminant migration through the capping layer or provide greater accumulation capacity, or reagents to encourage degradation or transformation of the contaminants.  
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{| class="wikitable mw-collapsible" style="float:left; margin-right:20px; text-align:center;"
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|+Table 1. Analyte list with acronyms and CAS numbers.
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|-
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!Compound
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! Acronym
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!CAS Number
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|-
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| 1,2-Dinitrobenzene (surrogate) ||'''1,2-DNB (surr.)''' || 528-29-0
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|-
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| 1,3-Dinitrobenzene || 1,3-DNB || 99-65-0
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|-
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| 1,3,5-Trinitrobenzene || 1,3,5-TNB || 99-35-4
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|-
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| 1,4-Dinitrobenzene || '''1,4-DNB (surr.)''' || 100-25-4
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|-
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| 2-Amino-4,6-dinitrotoluene || 2-Am-4,6-DNT || 35572-78-2
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|-
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| 2-Nitrophenol || '''2-NP''' || 88-75-5
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|-
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| 2-Nitrotoluene || 2-NT || 88-72-2
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|-
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| 2,4-Dinitrophenol || '''2,4-DNP''' || 51-28-5
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|-
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| 2,4-Dinitrotoluene || 2,4-DNT || 121-14-2
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|-
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| 2,4,6-Trinitrophenol || '''Picric Acid (PA)''' || 88-89-1
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|-
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| 2,4,6-Trinitrotoluene || 2,4,6-TNT || 118-96-7
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|-
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| 2,6-Dinitrotoluene || 2,6-DNT || 606-20-2
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|-
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| 3-Nitrotoluene || 3-NT || 99-08-1
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|-
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| 3,5-Dinitroaniline || 3,5-DNA || 618-87-1
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|-
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| 4-Amino-2,6-dinitrotoluene || 4-Am-2,6-DNT || 19406-51-0
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|-
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| 4-Nitrophenol || '''4-NP''' || 100-02-7
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|-
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| 4-Nitrotoluene || 4-NT || 99-99-0
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|-
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| 2,4-Dinitroanisole || '''DNAN''' || 119-27-7
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|-
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| Octahydro-1,3,5,7-tetranitro-1,3,5,7-tetrazocine || HMX || 2691-41-0
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|-
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| Nitrobenzene || NB || 98-95-3
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|-
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| Nitroglycerine || NG || 55-63-0
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|-
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| Nitroguanidine || '''NQ''' || 556-88-7
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|-
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| 3-Nitro-1,2,4-triazol-5-one || '''NTO''' || 932-64-9
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|-
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| ''ortho''-Nitrobenzoic acid || '''''o''-NBA (surr.)''' || 552-16-9
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|-
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| Pentaerythritol tetranitrate || PETN || 78-11-5
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|-
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| Hexahydro-1,3,5-trinitro-1,3,5-triazine || RDX || 121-82-4
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|-
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| N-Methyl-N-(2,4,6-trinitrophenyl)nitramide || Tetryl || 479-45-8
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|-
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| colspan="3" style="background-color:white;" | Note: Analytes in '''bold''' are not identified by EPA Method 8330B.
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|}
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[[File: ScircleFig1.png | thumb | 400px | Figure 1. Primary Method labeled chromatograms]]
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[[File: ScircleFig2.png | thumb | 400px | Figure 2. Secondary Method labeled chromatograms]]
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The&nbsp;primary&nbsp;intention of the analytical methods presented here is to support the monitoring of legacy and insensitive munitions contamination on test and training ranges, however legacy and insensitive munitions often accompany each other at demilitarization facilities, manufacturing facilities, and other environmental sites. Energetic materials typically appear on ranges as small, solid particulates and due to their varying functional groups and polarities, can partition in various environmental compartments<ref>Walsh, M.R., Temple, T., Bigl, M.F., Tshabalala, S.F., Mai, N. and Ladyman, M., 2017. Investigation of Energetic Particle Distribution from High‐Order Detonations of Munitions. Propellants, Explosives, Pyrotechnics, 42(8), pp. 932-941. [https://doi.org/10.1002/prep.201700089 doi: 10.1002/prep.201700089]</ref>. To ensure that contaminants are monitored and controlled at these sites and to sustainably manage them a variety of sample matrices (surface or groundwater, process waters, soil, and tissues) must be considered. (Process water refers to water used during industrial manufacturing or processing of legacy and insensitive munitions.) Furthermore, additional analytes must be added to existing methodologies as the usage of IM compounds changes and as new degradation compounds are identified.  Of note, relatively new IM formulations containing [[Wikipedia: Nitrotriazolone | NTO]], [[Wikipedia: 2,4-Dinitroanisole | DNAN]], and [[Wikipedia: Nitroguanidine | NQ]] are seeing use in [[Wikipedia: IMX-101 | IMX-101]], IMX-104, Pax-21 and Pax-41 (Table 1)<ref>Mainiero, C. 2015. Picatinny Employees Recognized for Insensitive Munitions. U.S. Army, Picatinny Arsenal Public Affairs.  [https://www.army.mil/article/148873/picatinny_employees_recognized_for_insensitive_munitions Open Access Press Release]</ref><ref>Frem, D., 2022. A Review on IMX-101 and IMX-104 Melt-Cast Explosives: Insensitive Formulations for the Next-Generation Munition Systems. Propellants, Explosives, Pyrotechnics, 48(1), e202100312. [https://doi.org/10.1002/prep.202100312 doi: 10.1002/prep.202100312]</ref>.
  
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The basic concept of a cap is illustrated in Figure 1. The Figure also illustrates that a cap is often a thin layer or layers relative to water depth and generally causes little disturbance to the underlying sediments or body of water in which it is placed.   Depending upon the erosive forces to which the cap may be subjected, the surface layer may be composed of relatively coarse material to withstand those erosive forces.  
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Sampling procedures for legacy and insensitive munitions are identical and utilize multi-increment sampling procedures found in USEPA Method 8330B Appendix A<ref name= "8330B"/>. Sample hold times, subsampling and quality control requirements are also unchanged. The key differences lie in the extraction methods and instrumental methods. Briefly, legacy munitions analysis of low concentration waters uses a single cartridge reverse phase [[Wikipedia: Solid-phase extraction | SPE]] procedure, and [[Wikipedia: Acetonitrile | acetonitrile]] (ACN) is used for both extraction and [[Wikipedia: Elution | elution]] for aqueous and solid samples<ref name= "8330B"/><ref>United States Environmental Protection Agency (USEPA), 2007. EPA Method 3535A (SW-846) Solid-Phase Extraction (SPE), Revision 1. [https://www.epa.gov/esam/epa-method-3535a-sw-846-solid-phase-extraction-spe USEPA Website]&nbsp; &nbsp;[[Media: epa-3535a.pdf | Method 3535A.pdf]]</ref>. An [[Wikipedia: High-performance_liquid_chromatography#Isocratic_and_gradient_elution | isocratic]] separation via reversed-phase C-18 column with 50:50 methanol:water mobile phase or a C-8 column with 15:85 isopropanol:water mobile phase is used to separate legacy munitions<ref name= "8330B"/>. While these procedures are sufficient for analysis of legacy munitions, alternative solvents, additional SPE cartridges, and a gradient elution are all required for the combined analysis of legacy and insensitive munitions.  
  
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Although a cap is typically thin compared to the water depth, it generally must be thicker than the biologically active zone (BAZ) of the sediments. The biologically active zone is that zone in which benthic organisms live and interact with the sediment.  Their activities tend to mix the BAZ (known as [[Wikipedia: Bioturbation | bioturbation]]) over the course of a few years and thus a cap that is thinner than the BAZ will tend to become intermixed with the underlying contaminated sediments.   Processes other than bioturbation including diffusion, advection or groundwater upwelling, hyporheic exchange near the interface, biogenic gas production and migration and underlying sediment consolidation can all lead to contaminant migration into and through a cap.  These occur at different rates and intensities and their assessment and evaluation ultimately governs the effectiveness of a cap and the feasibility of its use as a sediment remediation technology for a particular site.
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Previously, analysis of legacy and insensitive munitions required multiple analytical techniques, however the methods presented here combine the two munitions categories resulting in an HPLC-UV method and accompanying extraction methods for a variety of common sample matrices. A secondary HPLC-UV method and a HPLC-MS method were also developed as confirmatory methods. The methods discussed in this article were validated extensively by single-blind round robin testing and subsequent statistical treatment as part of ESTCP [https://serdp-estcp.mil/projects/details/d05c1982-bbfa-42f8-811d-51b540d7ebda ER19-5078]. Wherever possible, the quality control criteria in the Department of Defense Quality Systems Manual for Environmental Laboratories were adhered to<ref>US Department of Defense and US Department of Energy, 2021. Consolidated Quality Systems Manual (QSM) for Environmental Laboratories, Version 5.4. 387 pages. [https://www.denix.osd.mil/edqw/denix-files/sites/43/2021/10/QSM-Version-5.4-FINAL.pdf Free Download]&nbsp; &nbsp;[[Media: QSM-Version-5.4.pdf | QSM Version 5.4.pdf]]</ref>. Analytes included in the methods presented here are found in Table&nbsp;1.
  
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In general, capping is an effective remedial technology for contaminants that are strongly associated with the sediment solids including hydrophobic organic compounds such as high molecular weight [[Polycyclic Aromatic Hydrocarbons (PAHs) | polycyclic aromatic hydrocarbons (PAHs)]], [[Wikipedia: Polychlorinated biphenyl | polychlorinated biphenyls (PCBs)]], [[Wikipedia: Dioxins and dioxin-like compounds | dioxins]] and [[Wikipedia: DDT | DDTx]], but also [[Metal and Metalloid Contaminants | heavy metals]]. Hydrophobic organic compounds tend to strongly associate with the organic fraction of sediments so organic rich sediments or the addition of organic phases to the capping material can be very effective at containing these contaminants. Many of the common heavy metals of concern, including cadmium, copper, nickel, zinc, lead and mercury, tend to be associated with insoluble sulfides under strongly reducing conditions.  Since oxygen penetration into a capping layer is typically limited to a few cm or less at the surface, a cap serves to drive the underlying contaminated sediment toward strongly reducing conditions and, particularly in marine and estuarine sediments, encourage sulfate reduction leading to the formation of these insoluble sulfides.  The low solubility of these sulfides encourages retention by a capping layer and makes the cap extremely effective as a remedial approach for sediments with elevated concentrations of heavy metals.
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The chromatograms produced by the primary and secondary HPLC-UV methods are shown in Figure 1 and Figure 2, respectively. Chromatograms for each detector wavelength used are shown (315, 254, and 210 nm).
  
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A variety of tools have been developed to evaluate the processes leading to sorption and retardation of contaminants as well as processes leading to contaminant migration and release. The original references quantifying contaminant behavior in a sediment cap were explored in a series of papers in the early 1990s<ref name="Wang1991">Wang, X.Q., Thibodeaux, L.J., Valsaraj, K.T. and Reible, D.D., 1991. Efficiency of Capping Contaminated Bed Sediments in Situ. 1. Laboratory-Scale Experiments on Diffusion-Adsorption in the Capping Layer. Environmental Science and Technology, 25(9), pp.1578-1584.  [https://doi.org/10.1021/es00021a008 DOI: 10.1021/es00021a008]</ref><ref name="Thoma1993">Thoma, G.J., Reible, D.D., Valsaraj, K.T. and Thibodeaux, L.J., 1993. Efficiency of Capping Contaminated Bed Sediments in Situ 2. Mathematics of Diffusion-Adsorption in the Capping Layer. Environmental Science and Technology, 27(12), pp.2412-2419.  [https://doi.org/10.1021/es00048a015 DOI: 10.1021/es00048a015]</ref>. Since that time, design tools have been continuously improved. [https://www.depts.ttu.edu/ceweb/research/reiblesgroup/downloads.php CapSim] is a commonly used and current tool developed by Dr. Reible and collaborators. This tool can evaluate contaminant release from uncapped, capped, and treated sediments for purposes of design and evaluation.  The model formulation and structure is described in Shen et al. 2018<ref name="Shin2018">Shen, X., Lampert, D., Ogle, S. and Reible, D., 2018. A software tool for simulating contaminant transport and remedial effectiveness in sediment environments. Environmental Modelling and Software, 109, pp. 104-113.  [https://doi.org/10.1016/j.envsoft.2018.08.014 DOI: 10.1016/j.envsoft.2018.08.014]</ref>. One common use of such a tool is to evaluate the effect of various cap materials and thicknesses on the performance of a cap.
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==Extraction Methods==
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[[File: ScircleFig3.PNG |thumb|400px|Figure 3. Triple cartridge SPE setup]]
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[[File: ScircleFig4.PNG |thumb|400px|Figure 4. A flow chart of the soil extraction procedure]]
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===High&nbsp;Concentration&nbsp;Waters (> 1 ppm)===
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Aqueous samples suspected to contain the compounds of interest at concentrations detectable without any extraction or pre-concentration are suitable for analysis by direct injection. The method deviates from USEPA Method 8330B by adding a pH adjustment and use of MeOH rather than ACN for dilution<ref name= "8330B"/>. The pH adjustment is needed to ensure method accuracy for ionic compounds (like NTO or PA) in basic samples. A solution of 1% HCl/MeOH is added to both acidify and dilute the samples to a final acid concentration of 0.5% (vol/vol) and a final solvent ratio of 1:1 MeOH/H<sub><small>2</small></sub>O. The direct injection samples are then ready for analysis.
  
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==Cap Design and Materials for Chemical Containment==
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===Low Concentration Waters (< 1 ppm)===
−
An inert material such as sand can be effective as a capping material where contaminants are strongly associated with solids and where the operative site specific transport mechanisms do not lead to rapid contaminant migration through such a material. Additional contaminant containment can often be achieved through the placement of clean sediment, e.g. dredged material from a nearby location.  Other materials as cap layers or amendments may be useful to address particularly mobile contaminants or when particular degradative mechanisms can be exploited. The Anacostia River was the site of a demonstration that first tested “active” or “amended” capping in the field<ref name="Reible2003">Reible, D., Constant, D.W., Roberts, K. and Zhu, Y., 2003. Active capping demonstration project in anacostia DC. In Second International Conference on the Remediation of Contaminated Sediments: October.  Free download available from: [https://www.researchgate.net/profile/Danny-Reible/publication/237747790_ACTIVE_CAPPING_DEMONSTRATION_PROJECT_IN_ANACOSTIA_DC/links/0c96053861030b7699000000/ACTIVE-CAPPING-DEMONSTRATION-PROJECT-IN-ANACOSTIA-DC.pdf ResearchGate]</ref><ref name="Reible2006">Reible, D., Lampert, D., Constant, D., Mutch Jr, R.D. and Zhu, Y., 2006. Active Capping Demonstration in the Anacostia River, Washington, DC. Remediation Journal: The Journal of Environmental Cleanup Costs, Technologies and Techniques, 17(1), pp. 39-53.  [https://doi.org/10.1002/rem.20111 DOI: 10.1002/rem.20111]  Free download available from: [https://www.academia.edu/download/44146457/Remediation_Journal_Paper_2006.pdf Academia.edu]</ref>. Amended caps are often the best option when groundwater upwelling or other advective processes promote significant mobility of contaminants and the addition of sorbents can slow that contaminant migration<ref name="Ghosh2011">Ghosh, U., Luthy, R.G., Cornelissen, G., Werner, D. and Menzie, C.A., 2011. In-situ Sorbent Amendments: A New Direction in Contaminated Sediment Management. Environmental Science and Technology, 45(4), pp. 1163-1168. [https://doi.org/10.1021/es102694h DOI: 10.1021/es102694h]  Open access article from: [https://pubs.acs.org/doi/pdf/10.1021/es102694h American Chemical Society]&nbsp;&nbsp; [[Media: Ghosh2011.pdf | Report.pdf]]</ref>. Although a variety of materials have been proposed for sediment caps, a far smaller number of options have been successfully employed in the field.  
+
Aqueous samples suspected to contain the compounds of interest at low concentrations require extraction and pre-concentration using solid phase extraction (SPE). The SPE setup described here uses a triple cartridge setup shown in Figure 3. Briefly, the extraction procedure loads analytes of interest onto the cartridges in this order: Strata<sup><small>TM</small></sup> X, Strata<sup><small>TM</small></sup> X-A, and Envi-Carb<sup><small>TM</small></sup>. Then the cartridge order is reversed, and analytes are eluted via a two-step elution, resulting in 2 extracts (which are combined prior to analysis). Five milliliters of MeOH is used for the first elution, while 5 mL of acidified MeOH (2% HCl) is used for the second elution. The particular SPE cartridges used are noncritical so long as cartridge chemistries are comparable to those above.  
−
+
 
−
Metals migration is very site dependent due to the potential for many metals to complex with other species in the interstitial water and the specific metal speciation present at a site. Often, the strongly reducing environment beneath a cap renders many common metals unavailable through the formation of metal sulfides. In such cases, a simple sand cap can be very effective. Amended caps to manage metal contaminated sediments may be advantageous when site specific conditions lead to elevated metals mobility, but should be supported with site specific testing<ref name="Viana2008">Viana, P.Z., Yin, K. and Rockne, K.J., 2008. Modeling Active Capping Efficacy. 1. Metal and Organometal Contaminated Sediment Remediation. Environmental Science and Technology, 42(23), pp. 8922-8929. [https://doi.org/10.1021/es800942t DOI: 10.1021/es800942t]</ref>.
+
{| class="wikitable mw-collapsible" style="float:left; margin-right:20px; text-align:center;"
 +
|+Table 2. Primary HPLC-UV mobile phase gradient method concentrations
 +
|-
 +
| colspan="5" style="background-color:white;"| Method run time = 48 minutes; Column temperature = 25&deg;C<br>Injection volume = 50 &mu;L; Flow rate = 1.0 mL/min<br>Detector wavelengths = 210, 254, and 310 nm
 +
|-
 +
! Time<br>(min)
 +
! Reagent Water<br>(%)
 +
! MeOH<br>(%)
 +
! 0.1% TFA/Water<br>(%)
 +
! ACN<br>(%)
 +
|-
 +
| 0.00 || 89 || 3 || 3 || 5
 +
|-
 +
| 2.00 || 89 || 3 || 3 || 5
 +
|-
 +
| 2.20 || 52 || 40 || 3 || 5
 +
|-
 +
| 12.5 || 52 || 40 || 3 || 5
 +
|-
 +
| 19.0 || 57 ||35 || 3 || 5
 +
|-
 +
| 28.0 || 48 || 44 || 3 || 5
 +
|-
 +
| 32.0 || 48 || 44 || 3 || 5
 +
|-
 +
| 44.0 || 32 || 60 || 3 || 5
 +
|-
 +
| 44.1 || 89 || 3 || 3 || 5
 +
|-
 +
| 48.0 || 89 || 3 || 3 || 5
 +
|}
  
−
For hydrophobic organic contaminants, cap amendments that directly control groundwater upwelling and also sorbents that can remove migrating contaminants from that groundwater have been successfully employed.  Examples include clay materials such as AquaBlok<sup>&reg;</sup> for permeability control, sorbents such as [[Wikipedia: Activated carbon | activated carbon]] for truly dissolved contaminants, and [[Wikipedia: Organoclay | organophilic clays]] for separate phase contaminants.
+
===Soils=== 
 +
Soil collection, storage, drying and grinding procedures are identical to the USEPA Method 8330B procedures<ref name= "8330B"/>; however, the solvent extraction procedure differs in the number of sonication steps, sample mass and solvent used. A flow chart of the soil extraction procedure is shown in Figure 4. Soil masses of approximately 2 g and a sample to solvent ratio of 1:5 (g/mL) are used for soil extraction. The extraction is carried out in a sonication bath chilled below 20 ⁰C and is a two-part extraction, first extracting in MeOH (6 hours) followed by a second sonication in 1:1 MeOH:H<sub><small>2</small></sub>O solution (14 hours). The extracts are centrifuged, and the supernatant is filtered through a 0.45 μm PTFE disk filter.  
  
−
The placement of clean sediment as an ''in situ'' cap can be difficult when the material is fine grained or has a low density.  Capping with a layer of coarse grained material such as clean sand mitigates this issue although clean sands have minimal sorption capacity.  Because of this limitation, sand caps may not be sufficient for achieving remedial goals in sites where contamination levels are high or transport rates are fast due to pore water upwelling or tidal pumping effects. Conditions such as these may require the use of “active” amendments to reduce transport rates.  
+
The solvent volume should generally be 10 mL but if different soil masses are required, solvent volume should be 5 mL/g. The extraction results in 2 separate extracts (MeOH and MeOH:H<sub><small>2</small></sub>O) that are combined prior to analysis.
−
   
 
−
Capping with clean sand provides a physical barrier between the underlying contaminated material and the overlying water, stabilizes the underlying sediment to prevent re-suspension of contaminated particles, and can reduce chemical exposure under certain conditions.  Sand primarily provides a passive barrier to the downward penetration of bioturbating organisms and the upward movement of sediment or contaminants.  Although conventional sandy caps can often be an effective means of managing contaminated sediments, there are conditions when sand caps may not be capable of achieving design objectives.  Some factors that reduce the effectiveness of sand caps include:
 
  
−
*erosion and loss of cap integrity
+
===Tissues===
−
*high groundwater upwelling rates
 
−
*mobile (low sorption) contaminants of concern (COCs)
 
−
*high COC concentrations
 
−
*unusually toxic COCs
 
−
*the presence of tidal influences
 
−
*the presence of non-aqueous phase liquids (NAPLs)
 
−
*high rates of gas ebullition
 
  
−
Of these, the first three are common limitations to capping and often control the ability to effectively design and implement a cap as a sediment remedial strategy. In these cases, it may be possible to offset these issues by increasing the thickness of the cap. However, the required thickness can reach infeasible levels in shallow streams or navigable water bodies.  In addition, increased construction costs associated with thick caps may become prohibitive.  As a result of these issues, caps that use alternative materials (also known as active caps) to reduce the thickness or increase the protectiveness of a cap may be necessary. The materials in active caps are designed to interact with the COCs to enhance the containment properties of the cap.  
+
Tissue matrices are extracted by 18-hour sonication using a ratio of 1 gram of wet tissue per 5 mL of MeOH. This extraction is performed in a sonication bath chilled below 20 ⁰C and the supernatant (MeOH) is filtered through a 0.45 μm PTFE disk filter.  
  
−
[[Wikipedia: Apatite | Apatites]] are a class of naturally occurring minerals that have been investigated as a sorbent for metals in soils and sediments<ref name="Melton2003">Melton, J.S., Crannell, B.S., Eighmy, T.T., Wilson, C. and Reible, D.D., 2003. Field Trial of the UNH Phosphate-Based Reactive Barrier Capping System for the Anacostia River. EPA Grant R819165-01-0</ref><ref name="Reible2003"/><ref name="Knox2012">Knox, A.S., Paller, M.H. and Roberts, J., 2012. Active Capping Technology—New Approaches for In Situ Remediation of Contaminated Sediments. Remediation Journal, 22(2), pp.93-117.  [https://doi.org/10.1002/rem.21313 DOI: 10.1002/rem.21313]  Free download available from: [https://www.researchgate.net/profile/Anna-Knox-2/publication/233374607_Active_Capping_Technology-New_Approaches_for_In_Situ_Remediation_of_Contaminated_Sediments/links/5a7de4c5aca272a73765c344/Active-Capping-Technology-New-Approaches-for-In-Situ-Remediation-of-Contaminated-Sediments.pdf ResearchGate]</ref>.  Apatites consist of a matrix of calcium phosphate and various other common anions, including fluoride, chloride, hydroxide, and occasionally carbonate. Metals are sequestered either through direct ion exchange with the calcium atom or dissolution of hydroxyapatite followed by precipitation of lead apatite.  [[Wikipedia: Zeolite | Zeolites]], which are microporous aluminosilicate minerals with a high cationic exchange capacity (CEC), have also been proposed to manage metal species<ref name="Zhan2019">Zhan, Y., Yu, Y., Lin, J., Wu, X., Wang, Y. and Zhao, Y., 2019. Simultaneous control of nitrogen and phosphorus release from sediments using iron-modified zeolite as capping and amendment materials. Journal of Environmental Management, 249, p.109369. [https://doi.org/10.1016/j.jenvman.2019.109369 DOI: 10.1016/j.jenvman.2019.109369]</ref>.
+
Due to the complexity of tissue matrices, an additional tissue cleanup step, adapted from prior research, can be used to reduce interferences<ref name="RussellEtAl2014">Russell, A.L., Seiter, J.M., Coleman, J.G., Winstead, B., Bednar, A.J., 2014. Analysis of munitions constituents in IMX formulations by HPLC and HPLC-MS. Talanta, 128, pp. 524–530. [https://doi.org/10.1016/j.talanta.2014.02.013 doi: 10.1016/j.talanta.2014.02.013]</ref><ref name="CrouchEtAl2020"/>. The cleanup procedure uses small scale chromatography columns prepared by loading 5 ¾” borosilicate pipettes with 0.2 g activated silica gel (100–200 mesh). The columns are wetted with 1 mL MeOH, which is allowed to fully elute and then discarded prior to loading with 1 mL of extract and collecting in a new amber vial. After the extract is loaded, a 1 mL aliquot of MeOH followed by a 1 mL aliquot of 2% HCL/MeOH is added. This results in a 3 mL silica treated tissue extract. This extract is vortexed and diluted to a final solvent ratio of 1:1 MeOH/H<sub><small>2</small></sub>O before analysis.
−
 
 
−
It is possible to create a hydrophobic, sorbing layer for non-polar organics by exchanging a cationic surfactant onto the surface of clays such as zeolites and bentonites,. Organoclay is a modified bentonite containing such substitutions that has been evaluated for control of non-aqueous phase NAPLs and other organic contaminants<ref name="Reible2007">Reible, D.D., Lu, X., Moretti, L., Galjour, J. and Ma, X., 2007. Organoclays for the capping of contaminated sediments. AIChE Annual Meeting. ISBN: 978-081691022-9</ref>.  An organoclay cap has been implemented for sediment remediation at the McCormick and Baxter site in Portland, OR<ref name="Parrett2005">Parrett, K. and Blishke, H., 2005. 23-Acre Multilayer Sediment Cap in Dynamic Riverine Environment Using Organoclay an Adsorptive Capping Material. Presentation to Society of Environmental Toxicology and Chemistry (SETAC), 26th Annual Meeting.</ref>.  A similar organic sorbing phase can be formed by treating zeolites with surfactants but this approach has not been reported for contaminated sediments.  
 
  
−
Activated carbon is a strong sorbent of hydrophobic organic compounds and has been used as a [[In Situ Treatment of Contaminated Sediments with Activated Carbon | treatment for sediments]] or as an active sorbent within a capping layer<ref name="Zimmerman2004">Zimmerman, J.R., Ghosh, U., Millward, R.N., Bridges, T.S. and Luthy, R.G., 2004. Addition of Carbon Sorbents to Reduce PCB and PAH Bioavailability in Marine Sediments: Physicochemical Tests. Environmental Science and Technology, 38(20), pp. 5458-5464.  [https://doi.org/10.1021/es034992v DOI: 10.1021/es034992v]</ref><ref name="Werner2005">Werner, D., Higgins, C.P. and Luthy, R.G., 2005. The sequestration of PCBs in Lake Hartwell sediment with activated carbon. Water Research, 39(10), pp. 2105-2113.  [https://doi.org/10.1016/j.watres.2005.03.019 DOI: 10.1016/j.watres.2005.03.019]</ref><ref name="Abel2018">Abel, S. and Akkanen, J., 2018. A Combined Field and Laboratory Study on Activated Carbon-Based Thin Layer Capping in a PCB-Contaminated Boreal Lake. Environmental Science and Technology, 52(8), pp. 4702-4710. [https://doi.org/10.1021/acs.est.7b05114 DOI: 10.1021/acs.est.7b05114] Open access article available from: [https://pubs.acs.org/doi/pdf/10.1021/acs.est.7b05114 American Chemical Society]&nbsp;&nbsp; [[Media: Abel2018.pdf | Report.pdf]]</ref><ref name="Payne 2018">Payne, R.B., Ghosh, U., May, H.D., Marshall, C.W. and Sowers, K.R., 2019. A Pilot-Scale Field Study: In Situ Treatment of PCB-Impacted Sediments with Bioamended Activated Carbon. Environmental Science and Technology, 53(5), pp. 2626-2634. [https://doi.org/10.1021/acs.est.8b05019 DOI: 10.1021/acs.est.8b05019]</ref><ref name="Yan2020">Yan, S., Rakowska, M., Shen, X., Himmer, T., Irvine, C., Zajac-Fay, R., Eby, J., Janda, D., Ohannessian, S. and Reible, D.D., 2020. Bioavailability Assessment in Activated Carbon Treated Coastal Sediment with In situ and Ex situ Porewater Measurements. Water Research, 185, p. 116259.  [https://doi.org/10.1016/j.watres.2020.116259 DOI: 10.1016/j.watres.2020.116259]</ref>.  Placement of activated carbon for sediment capping is difficult due to the near neutral buoyancy of the material but it has been applied in this manner in relatively low energy environments such as Onondaga Lake, Syracuse, NY<ref name="Vlassopoulos2017">Vlassopoulos, D., Russell, K., Larosa, P., Brown, R., Mohan, R., Glaza, E., Drachenberg, T., Reible, D., Hague, W., McAuliffe, J. and Miller, S., 2017. Evaluation, Design, and Construction of Amended Reactive Caps to Restore Onondaga Lake, Syracuse, New York, USA. Journal of Marine Environmental Engineering, 10(1), pp. 13-27.  Free download available from: [https://www.researchgate.net/publication/317762995_Evaluation_design_and_construction_of_amended_reactive_caps_to_restore_Onondaga_lake_Syracuse_New_York_USA ResearchGate]</ref>.  Alternatives in higher energy environments include placement of activated carbon in a mat such as the CETCO Reactive Core Mat (RCM)<sup>&reg;</sup> or Huesker Tektoseal<sup>&reg;</sup>, or as a composite material such as SediMite<sup>&reg;</sup> or AquaGate<sup>&reg;</sup>.  In the case of the mats, powdered or granular activated carbon can be placed in a controlled layer while the density of the composite materials is such that they can be broadcast from the surface and allowed to settle to the bottom.  In a sediment treatment application, the composite material would either be worked into the surface or allowed to intermix gradually by bioturbation and other processes.  In a capping application, the mat or composite material would typically be combined or overlain with a sand or other capping layer to keep it in place and to provide a chemical isolation layer away from the sediment surface.  
+
==HPLC-UV and HPLC-MS Methods==
 +
{| class="wikitable mw-collapsible" style="float:left; margin-right:20px; text-align:center;"
 +
|+Table 3. Secondary HPLC-UV mobile phase gradient method concentrations
 +
|-
 +
| colspan="5" style="background-color:white;" | Method run time = 43 minutes; Column temperature = 25&deg;C<br>Injection volume = 50 &mu;L; Flow rate = 0.8 mL/min<br>Detector wavelengths = 210, 254, and 310 nm
 +
|-
 +
! Time<br>(min)
 +
! Reagent Water<br>(%)
 +
! MeOH<br>(%)
 +
! 0.1% TFA/Water<br>(%)
 +
! ACN<br>(%)
 +
|-
 +
| 0.00 || 75 || 10 || 10 || 5
 +
|-
 +
| 2.50 || 75 || 10 || 10 || 5
 +
|-
 +
| 2.60 || 39 || 46 ||10 || 5
 +
|-
 +
| 9.00 || 39 || 46 ||10 || 5
 +
|-
 +
| 9.10 || 33.5 || 51.5 || 10 || 5
 +
|-
 +
| 15.00 || 35 || 50 || 10 || 5
 +
|-
 +
| 15.10 || 43 || 42 || 10 || 5
 +
|-
 +
| 33.00 || 30 || 55 || 10 || 5
 +
|-
 +
| 33.10 || 75 || 10 || 10 || 5
 +
|-
 +
| 43.00 || 75 || 10 || 10 || 5
 +
|}
 +
{| class="wikitable mw-collapsible" style="float:right; margin-left:20px; text-align:center;"
 +
|+Table 4. Ionization source and detector parameters
 +
|-
 +
! Parameter
 +
! Value
 +
|-
 +
| Ionization Source || APCI
 +
|-
 +
| Ionization Mode || Negative
 +
|-
 +
| Drying Gas Temperature (&deg;C) || 350
 +
|-
 +
| Vaporizer Temperature (&deg;C) || 325
 +
|-
 +
| Drying Gas Flow (L/min) || 4.0
 +
|-
 +
| Nebulizer Pressure (psig) || 40
 +
|-
 +
| Corona Current (&mu;A) || 10
 +
|-
 +
| Capillary Potential (V) || 1500
 +
|-
 +
| Mass Range || 40 – 400
 +
|-
 +
| Fragmentor || 100
 +
|-
 +
| Gain || 1
 +
|-
 +
| Threshold || 0
 +
|-
 +
| Step Size || 0.20
 +
|-
 +
| Speed (&mu;/sec) || 743
 +
|-
 +
| Peak Width (min) || 0.06
 +
|-
 +
| Cycle Time (sec/cycle) || 0.57
 +
|}
  
−
As an alternative to a sorptive capping amendment, low-permeability cap amendments have been proposed to enhance cap design life by decreasing pore water advection. Low permeability clays are an effective means to divert upwelling groundwater away from a contaminated sediment area but are difficult to place in the aqueous environment.  Bentonite clays can be placed in mats similar to what is done to provide a low permeability liner in landfills. There are also commercial products that can place clays directly such as the composite material AquaBlok<sup>&reg;</sup>, a bentonite clay and polymer based mineral around an aggregate core<ref name="Barth2008">Barth, E.F., Reible, D. and Bullard, A., 2008. Evaluation of the physical stability, groundwater seepage control, and faunal changes associated with an AquaBlok<sup>&reg;</sup> sediment cap. Remediation: The Journal of Environmental Cleanup Costs, Technologies and Techniques, 18(4), pp.63-70.  [https://doi.org/10.1002/rem.20183 DOI: 10.1002/rem.20183]</ref>.
+
The Primary HPLC-UV method uses a Phenomenex Synergi 4 µm Hydro-RP column (80Å, 250 x 4.6 mm), or comparable, and is based on both the HPLC method found in USEPA 8330B and previous work<ref name= "8330B"/><ref name="RussellEtAl2014"/><ref name="CrouchEtAl2020"/>. This separation relies on a reverse phase column and uses a gradient elution, shown in Table 2. Depending on the analyst’s needs and equipment availability, the method has been proven to work with either 0.1% TFA or 0.25% FA (vol/vol) mobile phase. Addition of a guard column like a Phenomenex SecurityGuard AQ C18 pre-column guard cartridge can be optionally used. These optional changes to the method have no impact on the method’s performance.  
−
 
 
−
Sediment caps become colonized by microorganisms from the sediments and surface water and potentially become a zone of pollutant biotransformation over time. Aerobic degradation occurs only near the solids-water interface in which benthic organisms are active and thus there might still be significant benthic organism exposure to contaminants. Biotransformation in the anaerobic zone of a cap, which typically extends well beyond the zone of benthic activity, could significantly reduce the risk of pollutant exposure but successful caps encouraging deep degradation processes have not been demonstrated beyond the laboratory.  The addition of materials such as nutrients and oxygen releasing compounds for enhancing the attenuation of contaminants through biodegradation has also been assessed but not applied in the field.  Short term improvements in biodegradation rates can be achieved through tailoring of conditions or addition of nutrients but long term efficacy has not been demonstrated<ref name="Pagnozzi2020">Pagnozzi, G., Carroll, S., Reible, D.D. and Millerick, K., 2020. Biological Natural Attenuation and Contaminant Oxidation in Sediment Caps: Recent Advances and Future Opportunities. Current Pollution Reports, pp.1-14. [https://doi.org/10.1007/s40726-020-00153-5 DOI: 10.1007/s40726-020-00153-5]</ref>. 
 
−
[[File: SedCapFig2.png | thumb |600px|Figure 2. A conceptualization of a cap with accompanying habitat layer]]
 
  
−
==Cap Design and Materials for Habitat Restoration==
+
The Secondary HPLC-UV method uses a Restek Pinnacle II Biphenyl 5 µm (150 x 4.6 mm) or comparable column and is intended as a confirmatory method. Like the Primary method, this method can use an optional guard column and utilizes a gradient elution, shown in Table 3.
−
In addition to providing chemical isolation and containment, a cap can also be used to provide improvements for organisms by enhancing the habitat characteristics of the bottom substrate<ref name="Yozzo2004">Yozzo, D.J., Wilber, P. and Will, R.J., 2004. Beneficial use of dredged material for habitat creation, enhancement, and restoration in New York–New Jersey Harbor. Journal of Environmental Management, 73(1), pp. 39-52.  [https://doi.org/10.1016/j.jenvman.2004.05.008 DOI: 10.1016/j.jenvman.2004.05.008]</ref><ref name="Zhang2016">Zhang, C., Zhu, M.Y., Zeng, G.M., Yu, Z.G., Cui, F., Yang, Z.Z. and Shen, L.Q., 2016. Active capping technology: a new environmental remediation of contaminated sediment. Environmental Science and Pollution Research, 23(5), pp.4370-4386.  [https://doi.org/10.1007/s11356-016-6076-8 DOI: 10.1007/s11356-016-6076-8]</ref><ref name="Vlassopoulos2017"/>. Often, contaminated sediment environments are degraded for a variety of reasons in addition to the toxic constituents.  One way to overcome this is to provide both a habitat layer and chemical isolation or contaminant capping layer. Figure 2 illustrates just such a design providing a more appropriate habitat enhancing substrate, in this case by incorporation additional organic material, vegetation and debris, which is often used by fish species for protection, into the surface layer. In a high energy environment, it should be recognized that it may not be possible to keep a suitable habitat layer in place during high flow events.  This would be true of suitable habitat that had developed naturally as well as a constructed habitat layer and it is presumed that if such a habitat is the normal condition of the waterbody that it will recover over time between such high flow events.
+
 +
For instruments equipped with a mass spectrometer (MS), a secondary MS method is available and was developed alongside the Primary UV method. The method was designed for use with a single quadrupole MS equipped with an atmospheric pressure chemical ionization (APCI) source, such as an Agilent 6120B. A majority of the analytes shown in Table 1 are amenable to this MS method, however nitroglycerine (which is covered extensively in USEPA method 8332) and 2-,3-, and 4-nitrotoluene compounds aren’t compatible with the MS method. MS method parameters are shown in Table 4.
  
 
==Summary==
 
==Summary==
−
Clean substrate can be placed at the sediment-water interface for the purposes of reducing exposure to and risk from contaminants in the sediments.  The cap can consist of simple materials such as sand designed to physically stabilize contaminated sediments and separate the benthic community from those contaminants or may include other materials designed to sequester contaminants even under adverse conditions including strong groundwater upwelling or highly mobile contaminants. The surface of a cap may be designed of coarse material such as gravel or cobble to be stable under high flow events or designed to be more appropriate habitat for benthic and aquatic organisms.  As a result of its flexibility, simplicity and low cost relative to its effectiveness, capping is one of the most prevalent remedial technologies for sediments.  
+
The extraction methods and instrumental methods in this article build upon prior munitions analytical methods by adding new compounds, combining legacy and insensitive munitions analysis, and expanding usable sample matrices. These methods have been verified through extensive round robin testing and validation, and while the methods are somewhat challenging, they are crucial when simultaneous analysis of both insensitive and legacy munitions is needed.  
  
 
==References==
 
==References==
Line 74: Line 249:
  
 
==See Also==
 
==See Also==
 +
*[[Media: ERDC_TR-21-12.pdf | Preparative, Extraction, and Analytical Methods for Simultaneous Determination of Legacy and Insensitive Munition (IM) Constituents in Aqueous, Soil or Sediment, and Tissue Matrices]]
 +
*[https://serdp-estcp.mil/focusareas/9f7a342a-1b13-4ce5-bda0-d7693cf2b82d/uxo#subtopics  SERDP/ESTCP Focus Areas – UXO – Munitions Constituents]
 +
*[https://denix.osd.mil/edqw/home/  Environmental Data Quality Workgroup]

Latest revision as of 21:59, 26 September 2024

Sediment Porewater Dialysis Passive Samplers for Inorganics (Peepers)

Sediment porewater dialysis passive samplers, also known as “peepers,” are sampling devices that allow the measurement of dissolved inorganic ions in the porewater of a saturated sediment. Peepers function by allowing freely-dissolved ions in sediment porewater to diffuse across a micro-porous membrane towards water contained in an isolated compartment that has been inserted into sediment. Once retrieved after a deployment period, the resulting sample obtained can provide concentrations of freely-dissolved inorganic constituents in sediment, which provides measurements that can be used for understanding contaminant fate and risk. Peepers can also be used in the same manner in surface water, although this article is focused on the use of peepers in sediment.

Related Article(s):


Contributor(s):

  • Florent Risacher, M.Sc.
  • Jason Conder, Ph.D.

Key Resource(s):

  • A review of peeper passive sampling approaches to measure the availability of inorganics in sediment porewater[1]
  • Best Practices User’s Guide: Standardizing Sediment Porewater Passive Samplers for Inorganic Constituents of Concern[2]

Introduction

Biologically available inorganic constituents associated with sediment toxicity can be quantified by measuring the freely-dissolved fraction of contaminants in the porewater[3][4]. Classical sediment porewater analysis usually consists of collecting large volumes of bulk sediments which are then mechanically squeezed or centrifuged to produce a supernatant, or suction of porewater from intact sediment, followed by filtration and collection[5]. The extraction and measurement processes present challenges due to the heterogeneity of sediments, physical disturbance, high reactivity of some complexes, and interaction between the solid and dissolved phases, which can impact the measured concentration of dissolved inorganics[6]. For example, sampling disturbance can affect redox conditions[7][8], which can lead to under or over representation of inorganic chemical concentrations relative to the true dissolved phase concentration in the sediment porewater[9][5].

To address the complications with mechanical porewater sampling, passive sampling approaches for inorganics have been developed to provide a method that has a low impact on the surrounding geochemistry of sediments and sediment porewater, thus enabling more precise measurements of inorganics[4]. Sediment porewater dialysis passive samplers, also known as “peepers,” were developed more than 45 years ago[10] and refinements to the method such as the use of reverse tracers have been made, improving the acceptance of the technology as decision-making tool.

Peeper Designs

Peepers (Figure 1) are inert containers with a small volume (typically 1-100 mL) of purified water (“peeper water”) capped with a semi-permeable membrane. Peepers can be manufactured in a wide variety of formats (Figure 2, Figure 3) and deployed in in various ways.

Two designs are commonly used for peepers. Frequently, the designs are close adaptations of the original multi-chamber Hesslein design[10] (Figure 2), which consists of an acrylic sampler body with multiple sample chambers machined into it. Peeper water inside the chambers is separated from the outside environment by a semi-permeable membrane, which is held in place by a top plate fixed to the sampler body using bolts or screws. An alternative design consists of single-chamber peepers constructed using a single sample vial with a membrane secured over the mouth of the vial, as shown in Figure 3, and applied in Teasdale et al.[7], Serbst et al.[11], Thomas and Arthur[12], Passeport et al.[13], and Risacher et al.[2]. The vial is filled with deionized water, and the membrane is held in place using the vial cap or an o-ring. Individual vials are either directly inserted into sediment or are incorporated into a support structure to allow multiple single-chamber peepers to be deployed at once over a given depth profile (Figure 3).


Table 1. Analyte list with acronyms and CAS numbers.
Compound Acronym CAS Number
1,2-Dinitrobenzene (surrogate) 1,2-DNB (surr.) 528-29-0
1,3-Dinitrobenzene 1,3-DNB 99-65-0
1,3,5-Trinitrobenzene 1,3,5-TNB 99-35-4
1,4-Dinitrobenzene 1,4-DNB (surr.) 100-25-4
2-Amino-4,6-dinitrotoluene 2-Am-4,6-DNT 35572-78-2
2-Nitrophenol 2-NP 88-75-5
2-Nitrotoluene 2-NT 88-72-2
2,4-Dinitrophenol 2,4-DNP 51-28-5
2,4-Dinitrotoluene 2,4-DNT 121-14-2
2,4,6-Trinitrophenol Picric Acid (PA) 88-89-1
2,4,6-Trinitrotoluene 2,4,6-TNT 118-96-7
2,6-Dinitrotoluene 2,6-DNT 606-20-2
3-Nitrotoluene 3-NT 99-08-1
3,5-Dinitroaniline 3,5-DNA 618-87-1
4-Amino-2,6-dinitrotoluene 4-Am-2,6-DNT 19406-51-0
4-Nitrophenol 4-NP 100-02-7
4-Nitrotoluene 4-NT 99-99-0
2,4-Dinitroanisole DNAN 119-27-7
Octahydro-1,3,5,7-tetranitro-1,3,5,7-tetrazocine HMX 2691-41-0
Nitrobenzene NB 98-95-3
Nitroglycerine NG 55-63-0
Nitroguanidine NQ 556-88-7
3-Nitro-1,2,4-triazol-5-one NTO 932-64-9
ortho-Nitrobenzoic acid o-NBA (surr.) 552-16-9
Pentaerythritol tetranitrate PETN 78-11-5
Hexahydro-1,3,5-trinitro-1,3,5-triazine RDX 121-82-4
N-Methyl-N-(2,4,6-trinitrophenyl)nitramide Tetryl 479-45-8
Note: Analytes in bold are not identified by EPA Method 8330B.
Figure 1. Primary Method labeled chromatograms
Figure 2. Secondary Method labeled chromatograms

The primary intention of the analytical methods presented here is to support the monitoring of legacy and insensitive munitions contamination on test and training ranges, however legacy and insensitive munitions often accompany each other at demilitarization facilities, manufacturing facilities, and other environmental sites. Energetic materials typically appear on ranges as small, solid particulates and due to their varying functional groups and polarities, can partition in various environmental compartments[14]. To ensure that contaminants are monitored and controlled at these sites and to sustainably manage them a variety of sample matrices (surface or groundwater, process waters, soil, and tissues) must be considered. (Process water refers to water used during industrial manufacturing or processing of legacy and insensitive munitions.) Furthermore, additional analytes must be added to existing methodologies as the usage of IM compounds changes and as new degradation compounds are identified. Of note, relatively new IM formulations containing NTO, DNAN, and NQ are seeing use in IMX-101, IMX-104, Pax-21 and Pax-41 (Table 1)[15][16].

Sampling procedures for legacy and insensitive munitions are identical and utilize multi-increment sampling procedures found in USEPA Method 8330B Appendix A[17]. Sample hold times, subsampling and quality control requirements are also unchanged. The key differences lie in the extraction methods and instrumental methods. Briefly, legacy munitions analysis of low concentration waters uses a single cartridge reverse phase SPE procedure, and acetonitrile (ACN) is used for both extraction and elution for aqueous and solid samples[17][18]. An isocratic separation via reversed-phase C-18 column with 50:50 methanol:water mobile phase or a C-8 column with 15:85 isopropanol:water mobile phase is used to separate legacy munitions[17]. While these procedures are sufficient for analysis of legacy munitions, alternative solvents, additional SPE cartridges, and a gradient elution are all required for the combined analysis of legacy and insensitive munitions.

Previously, analysis of legacy and insensitive munitions required multiple analytical techniques, however the methods presented here combine the two munitions categories resulting in an HPLC-UV method and accompanying extraction methods for a variety of common sample matrices. A secondary HPLC-UV method and a HPLC-MS method were also developed as confirmatory methods. The methods discussed in this article were validated extensively by single-blind round robin testing and subsequent statistical treatment as part of ESTCP ER19-5078. Wherever possible, the quality control criteria in the Department of Defense Quality Systems Manual for Environmental Laboratories were adhered to[19]. Analytes included in the methods presented here are found in Table 1.

The chromatograms produced by the primary and secondary HPLC-UV methods are shown in Figure 1 and Figure 2, respectively. Chromatograms for each detector wavelength used are shown (315, 254, and 210 nm).

Extraction Methods

Figure 3. Triple cartridge SPE setup
Figure 4. A flow chart of the soil extraction procedure

High Concentration Waters (> 1 ppm)

Aqueous samples suspected to contain the compounds of interest at concentrations detectable without any extraction or pre-concentration are suitable for analysis by direct injection. The method deviates from USEPA Method 8330B by adding a pH adjustment and use of MeOH rather than ACN for dilution[17]. The pH adjustment is needed to ensure method accuracy for ionic compounds (like NTO or PA) in basic samples. A solution of 1% HCl/MeOH is added to both acidify and dilute the samples to a final acid concentration of 0.5% (vol/vol) and a final solvent ratio of 1:1 MeOH/H2O. The direct injection samples are then ready for analysis.

Low Concentration Waters (< 1 ppm)

Aqueous samples suspected to contain the compounds of interest at low concentrations require extraction and pre-concentration using solid phase extraction (SPE). The SPE setup described here uses a triple cartridge setup shown in Figure 3. Briefly, the extraction procedure loads analytes of interest onto the cartridges in this order: StrataTM X, StrataTM X-A, and Envi-CarbTM. Then the cartridge order is reversed, and analytes are eluted via a two-step elution, resulting in 2 extracts (which are combined prior to analysis). Five milliliters of MeOH is used for the first elution, while 5 mL of acidified MeOH (2% HCl) is used for the second elution. The particular SPE cartridges used are noncritical so long as cartridge chemistries are comparable to those above.

Table 2. Primary HPLC-UV mobile phase gradient method concentrations
Method run time = 48 minutes; Column temperature = 25°C
Injection volume = 50 μL; Flow rate = 1.0 mL/min
Detector wavelengths = 210, 254, and 310 nm
Time
(min) 		Reagent Water
(%) 		MeOH
(%) 		0.1% TFA/Water
(%) 		ACN
(%)
0.00 89 3 3 5
2.00 89 3 3 5
2.20 52 40 3 5
12.5 52 40 3 5
19.0 57 35 3 5
28.0 48 44 3 5
32.0 48 44 3 5
44.0 32 60 3 5
44.1 89 3 3 5
48.0 89 3 3 5

Soils

Soil collection, storage, drying and grinding procedures are identical to the USEPA Method 8330B procedures[17]; however, the solvent extraction procedure differs in the number of sonication steps, sample mass and solvent used. A flow chart of the soil extraction procedure is shown in Figure 4. Soil masses of approximately 2 g and a sample to solvent ratio of 1:5 (g/mL) are used for soil extraction. The extraction is carried out in a sonication bath chilled below 20 ⁰C and is a two-part extraction, first extracting in MeOH (6 hours) followed by a second sonication in 1:1 MeOH:H2O solution (14 hours). The extracts are centrifuged, and the supernatant is filtered through a 0.45 μm PTFE disk filter.

The solvent volume should generally be 10 mL but if different soil masses are required, solvent volume should be 5 mL/g. The extraction results in 2 separate extracts (MeOH and MeOH:H2O) that are combined prior to analysis.

Tissues

Tissue matrices are extracted by 18-hour sonication using a ratio of 1 gram of wet tissue per 5 mL of MeOH. This extraction is performed in a sonication bath chilled below 20 ⁰C and the supernatant (MeOH) is filtered through a 0.45 μm PTFE disk filter.

Due to the complexity of tissue matrices, an additional tissue cleanup step, adapted from prior research, can be used to reduce interferences[20][21]. The cleanup procedure uses small scale chromatography columns prepared by loading 5 ¾” borosilicate pipettes with 0.2 g activated silica gel (100–200 mesh). The columns are wetted with 1 mL MeOH, which is allowed to fully elute and then discarded prior to loading with 1 mL of extract and collecting in a new amber vial. After the extract is loaded, a 1 mL aliquot of MeOH followed by a 1 mL aliquot of 2% HCL/MeOH is added. This results in a 3 mL silica treated tissue extract. This extract is vortexed and diluted to a final solvent ratio of 1:1 MeOH/H2O before analysis.

HPLC-UV and HPLC-MS Methods

Table 3. Secondary HPLC-UV mobile phase gradient method concentrations
Method run time = 43 minutes; Column temperature = 25°C
Injection volume = 50 μL; Flow rate = 0.8 mL/min
Detector wavelengths = 210, 254, and 310 nm
Time
(min) 		Reagent Water
(%) 		MeOH
(%) 		0.1% TFA/Water
(%) 		ACN
(%)
0.00 75 10 10 5
2.50 75 10 10 5
2.60 39 46 10 5
9.00 39 46 10 5
9.10 33.5 51.5 10 5
15.00 35 50 10 5
15.10 43 42 10 5
33.00 30 55 10 5
33.10 75 10 10 5
43.00 75 10 10 5
Table 4. Ionization source and detector parameters
Parameter Value
Ionization Source APCI
Ionization Mode Negative
Drying Gas Temperature (°C) 350
Vaporizer Temperature (°C) 325
Drying Gas Flow (L/min) 4.0
Nebulizer Pressure (psig) 40
Corona Current (μA) 10
Capillary Potential (V) 1500
Mass Range 40 – 400
Fragmentor 100
Gain 1
Threshold 0
Step Size 0.20
Speed (μ/sec) 743
Peak Width (min) 0.06
Cycle Time (sec/cycle) 0.57

The Primary HPLC-UV method uses a Phenomenex Synergi 4 µm Hydro-RP column (80Å, 250 x 4.6 mm), or comparable, and is based on both the HPLC method found in USEPA 8330B and previous work[17][20][21]. This separation relies on a reverse phase column and uses a gradient elution, shown in Table 2. Depending on the analyst’s needs and equipment availability, the method has been proven to work with either 0.1% TFA or 0.25% FA (vol/vol) mobile phase. Addition of a guard column like a Phenomenex SecurityGuard AQ C18 pre-column guard cartridge can be optionally used. These optional changes to the method have no impact on the method’s performance.

The Secondary HPLC-UV method uses a Restek Pinnacle II Biphenyl 5 µm (150 x 4.6 mm) or comparable column and is intended as a confirmatory method. Like the Primary method, this method can use an optional guard column and utilizes a gradient elution, shown in Table 3.

For instruments equipped with a mass spectrometer (MS), a secondary MS method is available and was developed alongside the Primary UV method. The method was designed for use with a single quadrupole MS equipped with an atmospheric pressure chemical ionization (APCI) source, such as an Agilent 6120B. A majority of the analytes shown in Table 1 are amenable to this MS method, however nitroglycerine (which is covered extensively in USEPA method 8332) and 2-,3-, and 4-nitrotoluene compounds aren’t compatible with the MS method. MS method parameters are shown in Table 4.

Summary

The extraction methods and instrumental methods in this article build upon prior munitions analytical methods by adding new compounds, combining legacy and insensitive munitions analysis, and expanding usable sample matrices. These methods have been verified through extensive round robin testing and validation, and while the methods are somewhat challenging, they are crucial when simultaneous analysis of both insensitive and legacy munitions is needed.

References

  1. ^ Risacher, F.F., Schneider, H., Drygiannaki, I., Conder, J., Pautler, B.G., and Jackson, A.W., 2023. A Review of Peeper Passive Sampling Approaches to Measure the Availability of Inorganics in Sediment Porewater. Environmental Pollution, 328, Article 121581. doi: 10.1016/j.envpol.2023.121581   Open Access Manuscript
  2. ^ 2.0 2.1 Risacher, F.F., Nichols, E., Schneider, H., Lawrence, M., Conder, J., Sweett, A., Pautler, B.G., Jackson, W.A., Rosen, G., 2023b. Best Practices User’s Guide: Standardizing Sediment Porewater Passive Samplers for Inorganic Constituents of Concern, ESTCP ER20-5261. Project Website   Report.pdf
  3. ^ Conder, J.M., Fuchsman, P.C., Grover, M.M., Magar, V.S., Henning, M.H., 2015. Critical review of mercury SQVs for the protection of benthic invertebrates. Environmental Toxicology and Chemistry, 34(1), pp. 6-21. doi: 10.1002/etc.2769   Open Access Article
  4. ^ 4.0 4.1 Cleveland, D., Brumbaugh, W.G., MacDonald, D.D., 2017. A comparison of four porewater sampling methods for metal mixtures and dissolved organic carbon and the implications for sediment toxicity evaluations. Environmental Toxicology and Chemistry, 36(11), pp. 2906-2915. doi: 10.1002/etc.3884
  5. ^ 5.0 5.1 Gruzalski, J.G., Markwiese, J.T., Carriker, N.E., Rogers, W.J., Vitale, R.J., Thal, D.I., 2016. Pore Water Collection, Analysis and Evolution: The Need for Standardization. In: Reviews of Environmental Contamination and Toxicology, Vol. 237, pp. 37–51. Springer. doi: 10.1007/978-3-319-23573-8_2
  6. ^ Peijnenburg, W.J.G.M., Teasdale, P.R., Reible, D., Mondon, J., Bennett, W.W., Campbell, P.G.C., 2014. Passive Sampling Methods for Contaminated Sediments: State of the Science for Metals. Integrated Environmental Assessment and Management, 10(2), pp. 179–196. doi: 10.1002/ieam.1502   Open Access Article
  7. ^ 7.0 7.1 Teasdale, P.R., Batley, G.E., Apte, S.C., Webster, I.T., 1995. Pore water sampling with sediment peepers. Trends in Analytical Chemistry, 14(6), pp. 250–256. doi: 10.1016/0165-9936(95)91617-2
  8. ^ Schroeder, H., Duester, L., Fabricius, A.L., Ecker, D., Breitung, V., Ternes, T.A., 2020. Sediment water (interface) mobility of metal(loid)s and nutrients under undisturbed conditions and during resuspension. Journal of Hazardous Materials, 394, Article 122543. doi: 10.1016/j.jhazmat.2020.122543   Open Access Article
  9. ^ Wise, D.E., 2009. Sampling techniques for sediment pore water in evaluation of reactive capping efficacy. Master of Science Thesis. University of New Hampshire Scholars’ Repository. 178 pages. Website   Report.pdf
  10. ^ 10.0 10.1 Hesslein, R.H., 1976. An in situ sampler for close interval pore water studies. Limnology and Oceanography, 21(6), pp. 912-914. doi: 10.4319/lo.1976.21.6.0912   Open Access Article
  11. ^ Serbst, J.R., Burgess, R.M., Kuhn, A., Edwards, P.A., Cantwell, M.G., Pelletier, M.C., Berry, W.J., 2003. Precision of dialysis (peeper) sampling of cadmium in marine sediment interstitial water. Archives of Environmental Contamination and Toxicology, 45(3), pp. 297–305. doi: 10.1007/s00244-003-0114-5
  12. ^ Thomas, B., Arthur, M.A., 2010. Correcting porewater concentration measurements from peepers: Application of a reverse tracer. Limnology and Oceanography: Methods, 8(8), pp. 403–413. doi: 10.4319/lom.2010.8.403   Open Access Article
  13. ^ Passeport, E., Landis, R., Lacrampe-Couloume, G., Lutz, E.J., Erin Mack, E., West, K., Morgan, S., Lollar, B.S., 2016. Sediment Monitored Natural Recovery Evidenced by Compound Specific Isotope Analysis and High-Resolution Pore Water Sampling. Environmental Science and Technology, 50(22), pp. 12197–12204. doi: 10.1021/acs.est.6b02961
  14. ^ Walsh, M.R., Temple, T., Bigl, M.F., Tshabalala, S.F., Mai, N. and Ladyman, M., 2017. Investigation of Energetic Particle Distribution from High‐Order Detonations of Munitions. Propellants, Explosives, Pyrotechnics, 42(8), pp. 932-941. doi: 10.1002/prep.201700089
  15. ^ Mainiero, C. 2015. Picatinny Employees Recognized for Insensitive Munitions. U.S. Army, Picatinny Arsenal Public Affairs. Open Access Press Release
  16. ^ Frem, D., 2022. A Review on IMX-101 and IMX-104 Melt-Cast Explosives: Insensitive Formulations for the Next-Generation Munition Systems. Propellants, Explosives, Pyrotechnics, 48(1), e202100312. doi: 10.1002/prep.202100312
  17. ^ 17.0 17.1 17.2 17.3 17.4 17.5 Cite error: Invalid <ref> tag; no text was provided for refs named 8330B
  18. ^ United States Environmental Protection Agency (USEPA), 2007. EPA Method 3535A (SW-846) Solid-Phase Extraction (SPE), Revision 1. USEPA Website    Method 3535A.pdf
  19. ^ US Department of Defense and US Department of Energy, 2021. Consolidated Quality Systems Manual (QSM) for Environmental Laboratories, Version 5.4. 387 pages. Free Download    QSM Version 5.4.pdf
  20. ^ 20.0 20.1 Russell, A.L., Seiter, J.M., Coleman, J.G., Winstead, B., Bednar, A.J., 2014. Analysis of munitions constituents in IMX formulations by HPLC and HPLC-MS. Talanta, 128, pp. 524–530. doi: 10.1016/j.talanta.2014.02.013
  21. ^ 21.0 21.1 Cite error: Invalid <ref> tag; no text was provided for refs named CrouchEtAl2020

See Also

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