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SPMD Technology Tutorial (3rd Edition)

(Updated January 03, 2002)

Prepared by
James N. Huckins
J.D. Petty
Jon A. Lebo
Carl E. Orazio
Randal C. Clark
Virginia L. Gibson

Table of Contents




Limitations of Conventional Sampling

SPMD Rationale and Design

Manufacturer or Source

Attributes and Applications

Performance Characteristics

Transport and Deployment

Sample Processing and Enrichment



Examples of Applications


Sampling Environmental Media

Residue Residence (Retention) Time

Appropriateness of Lipid Normalization

Further Information

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We gratefully acknowledge the financial support of the Biological Resources Division of the U.S. Geological Survey, U.S. Fish and Wildlife Service, the National Fish and Wildlife Foundation, National Science Foundation, the American Petroleum Institute, Chevron Oil Co., AMOCO Corp., and the Department of Defense.   We also appreciate the valuable contributions of Harry Prest, Robert Gale, Kees Booij, John Meadows, Kathy Echols, Tom Johnson, Don Tillitt, and Bruce Moring.

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Researchers at the U.S. Geological Survey's Columbia Environmental Research Center (CERC) developed and patented (government) a technology that has diverse potential environmental and industrial applications.  The technology is based on several different configurations of semipermeable membrane devices (SPMDs).  As passive in situ devices, SPMDs can be used for contaminant monitoring, exposure and toxicity assessment, and can also be used for analytical separations.  Although SPMD technology is becoming widely accepted, the literature provides little guidance on using SPMDs.  This document emphasizes the in situ concentration of contaminants for exposure and toxicity assessments and provides a brief tutorial for scientists.

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Semipermeable Membrane Devices (SPMDs)

Lipid-containing SPMDs represent an innovative passive sampling technology for monitoring and assessing trace levels of hydrophobic organic contaminants.  The SPMD is typically constructed from barefoot (no additives) layflat tubing of low-density polyethylene (LDPE).  The thin-walled (<100 µm) LDPE tubing used in SPMDs is normally described as nonporous.  However, random thermal motions of the polymer chains form transient cavities with maximum diameters of approximately 10 Å.  Because these cavities are extremely small and dynamic, hydrophobic solutes are essentially solubilized by the polymer.  The cross-sectional diameters of nearly all environmental contaminants are only slightly smaller than the polymeric cavities.  Therefore, only dissolved (i.e., readily bioavailable) organic contaminants diffuse through the membrane and are concentrated through time.  The sequestration media consist of both the thin film/plug of a large molecular weight (> 600 daltons) neutral lipid such as triolein and the LDPE membrane.  Contaminant residues concentrated in SPMDs are simultaneously recovered and separated from the lipid in intact SPMDs (after carefully cleaning exterior surface of the membrane) by dialysis in an organic solvent.

SPMDs accomplish three tasks simultaneously:


·        Mimics the bioconcentration of organic contaminants in fatty tissues of organisms


·        Provides a highly reproducible passive in situ sampler for monitoring contaminant levels, which is largely unaffected by many environmental stressors that affect biomonitoring organisms


·        Enables in situ concentration of trace organic contaminant mixtures for toxicity assessments and toxicity identification evaluation (TIE)

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Limitations of Conventional Sampling Methods

1.        Analysis of excised water and air samples reflects residue composition only at the moment of sampling and may fail to detect episodic contamination

2.        Quality control and physical difficulties are often encountered when large volumes of water and air must be collected and extracted for quantifying and assessing trace organic contaminants

3.        Concentrations of truly dissolved or readily bioavailable contaminants are not accurately measured by most conventional approaches

4.        Aquatic toxicity data, and threshold limit values for airborne exposures are based on dissolved or vapor phase concentrations, not total residue levels

5.        Standard low volume (< 4 L) techniques often fail to detect trace levels of bioconcentratable contaminants and seldom recover enough residue mass for bioassays

6.        Biomonitoring organisms may not accurately reflect environmental contaminant concentrations, because of residue metabolism/depuration and the effects of environmental stressors on organism health

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SPMD Approach to Contaminant Monitoring

Although passive diffusional monitors are widely accepted as the best method for determining occupational exposure of workers to ambient organic vapors, this type of integrative, or time-weighted average (TWA) approach has seldom been applied to concentrating / measuring trace contaminants in aquatic environments.  Passive integrative samplers act as infinite sinks for accumulated residues, i.e., no significant losses of sequestered residues occur even when ambient chemical concentrations fall during part of an exposure.  The following characteristics apply to SPMDs:

1.        The SPMD uptake of contaminants with log KOWs > 4.9 is usually integrative or linear (equilibrium not approached) during exposures £ 30 days, and SPMD concentrations are proportional to ambient environmental concentrations

2.        For organic compounds with log KOWS < 4.9, equilibrium concentrations of analytes may be reached or approached in < 30 days, but SPMD levels are still proportional to those in the sampled medium

3.        Integrative sampling reduces the probability of false-negatives, i.e., residues sequestered from episodic events are retained and thus can be detected

4.        Laboratory calibration (standard SPMD configuration) is required for estimating TWA ambient chemical concentrations from SPMD levels and these data are already available for several chemical classes

5.        Only dissolved or vapor phase (readily bioavailable) organic pollutants diffuse through or are sampled by the polymeric films or tubes used in SPMDs (e.g., LDPE and Silastic®)

6.        SPMDs mimic key mechanisms in contaminant bioconcentration; these include (a) the passive processes of contaminant diffusion through biomembranes (primarily the gill epithelium in fish) and (b) partitioning between an organism's lipids and the surrounding medium 

7.        The triolein used in SPMDs is a neutral lipid found in most aquatic organisms

8.        Because SPMDs are constructed of synthetic materials, they are more uniform and sample more reproducibly than biomonitoring organisms

9.        Although a combination of laboratory calibration data and the use of permeability/performance reference compounds (PRCs, see subsequent description of these QC standards) in deployed SPMDs are usually required for accurate estimates of contaminant concentrations in ambient media, the necessary information is available in several recent publications cited herein

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General Specifications of SPMDs


Thin-walled (50-100 µm) nonporous polymer film or tubes consisting of  


·        LDPE (low density polyethylene) 


·        Silicone or Silastic (option of plasma-treated surface)


·        Polypropylene


·        Ethylene vinyl acetate


·        Others

Potential sequestration phases:

Large molecular weight (» 600 daltons) nonpolar liquids or fluids such as


·        Neutral lipids


·        Silicone fluids


·        Other lipid-like organic fluids


·        The LDPE membrane alone

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Standard SPMD Configuration (Commercially Available)


LDPE layflat tubing manufactured without additives

Sequestration phases:

High-purity synthetic triolein (> 95%) and the LDPE membrane 

Dimensions (specifications):

Generally 2.5-cm wide (layflat) by 91.4-cm-long LDPE tubes (70-95 µm wall thickness and surface area is » 450 cm² or » 100 cm²/g SPMD) containing 1 mL (0.915 g) of triolein as a thin film.  Other lengths and widths can be used if the lipid-to-membrane mass ratio is maintained at » 0.2 and the membrane thickness is within the above range.

Note:  Nearly all SPMD calibration data are based on the standard SPMD.

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Commercial Availability

SPMD technology is the subject of two government patents (Huckins et al., U.S. Patents, #5,098,573 and #5,395,426) that were licensed to Environmental Sampling Technologies (EST), a division of Custom Industrial Analysis Labs, 1717 Commercial Drive, St. Joseph, MO 64503.  The patents cover both assembly of SPMD and dialytic recovery of analytes from SPMDs.  Various SPMD configurations and deployment apparatuses are available from the manufacturer.  The European source of SPMDs is ORIGO Hb, Trehorningen 34, S-922 66 Travelsjo, Sweden.

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SPMD Assembly and Quality Control Considerations

Assembling SPMDs requires considerable care to ensure that the finished product is free of contaminants. For sampling trace contaminants, the following quality control procedures are adhered to during SPMD manufacture:

1.        Synthetic triolein or lipid is used.  All new lots or batches are analyzed for contaminants, ampulated, and stored in a freezer until use

2.        A micropipettor is equipped with a total displacement plunger to accurately deliver small volumes of triolein

3.        Just before use in SPMD construction, SPMD tubing is batch-extracted with nanograde hexane or cyclohexane 

4.        To enclose triolein in SPMD layflat tubing, a heat sealer (e.g., the "Seal a Meal®" bag sealer) is used to create a molecular weld

5.        Because SPMDs are extremely efficient air samplers (PCB sampling rates for a 1-g triolein SPMD often approach and can exceed 10 m3/day), all assembly operations are conducted in an environmentally controlled chamber (clean room) with both vapor and particulate phase filters

6.        After assembly, SPMDs are sealed in gas-tight clean paint cans (solvent rinsed to remove cutting oils) for transport to deployment site

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·        Non-mechanical, passive device which is easy to deploy and requires no maintenance


·        Mimics uptake of dissolved contaminants by biota, yet precision of concentration data is greater


·        SPMD matrices can conveniently be cleaned up prior to use, while extensive depuration periods may be required to reduce contaminant levels in the tissues of biomonitoring organisms


·        Readily concentrates contaminant residues such as PAHs that are metabolized by many aquatic organisms


·        Once prepared, SPMDs can be stored frozen until the most appropriate deployment time, while biomonitoring organisms require care and feeding, and may be subject to seasonal availability


·        Analytical cleanup of exposed SPMDs is generally less difficult than biomonitoring organism tissues or sediment samples

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·        Determination of pollutant sources and relative levels


·        Detection of episodic chemical releases


·        Measurement of TWA concentrations of dissolved or vapor phase chemical concentrations


·        Determination of the readily bioavailable fraction (dissolved or vapor phase) of a chemical in an environmental compartment for predicting transport, fate, and residue toxicity


·        Estimation of organism exposure and bioconcentration


·        In situ biomimetic extraction of environmental contaminants for bioassay and immunoassay


·        Dialytic separations (SPMD membrane) of target analyte interferences in various matrices


·        Tool for TIE procedures

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Specifics of SPMD Sampling


·        Using the standard SPMD configuration with the LDPE membrane, only nonionic compounds are extracted or sampled because of the lack of membrane permeability by charged species


·        In general, molecules of organic compounds must be dissolved or in the vapor phase to be sampled by the LDPE SPMD membrane


·        SPMD linear uptake rates are expressed as the daily volume of water or air cleared of chemical by an SPMD, given in units of L or mLd-1, L or mLd-1g-1 and m³ d-1 g-1, respectively, and are independent of concentration


·        Theory and experimental data suggest that under relatively quiescent conditions (< 1 cm/sec, flow velocity at membrane surface) and moderate temperatures (i.e., 18-26ºC), the aqueous sampling rate of a 4.5 g standard SPMD (defined earlier) for most hydrophobic compounds ranges from about 0.5 to 10 L/d


·        Sampling rates can be affected by the physicochemical properties of the target compound (i.e., octanol-water or -air partition coefficient [KOW or KOA], polarity, molecular size/weight, and volatility) and environmental conditions of the exposure site (i.e., temperature, flow/turbulence, and biofouling level or the growth of a biofilm on the exterior membrane surface)


·        The driving force for uptake is directly related to the magnitude of a chemical's KOW/KOA or more specifically the KSPMD (equilibrium SPMD-water or -air partition coefficient) and for hydrophobic compounds the overall resistance to chemical uptake or mass transfer is inversely proportional to medium flow/turbulence, and directly proportional to the thickness of the aqueous boundary layer and the membrane (only when uptake controlled by membrane)


·        For exposure conditions of low to moderate flow/turbulence, SPMD uptake is under membrane control for compounds with log KOWs < 4.4 and under aqueous boundary layer control for compounds with log KOWs > 4.4


·        Sampling rates are unaffected by aqueous flow/turbulence only when a chemical is completely under membrane control


·        Temperature affects sampling rate, regardless of which step in the uptake process is rate limiting or has the most resistance to mass transfer


·        Given the wide range environmental exposure conditions, flow/turbulence effects are generally expected to have a greater impact on SPMD sampling rate than temperature or biofouling


·        The capacity or the total volume of water or air extracted by an SPMD at equilibrium is given by KSPMD MSPMD (where KSPMD is expressed as L, mL or cm³/g and MSPMD is the mass of the whole SPMD in g) and is roughly equivalent to KOW or KOA times MSPMD


·        The KOW or KOA can be viewed as the approximate volume (mL, cm³, or L) of water or air extracted by one g of SPMD at equilibrium


·        Depending on a compound’s KOW, residue uptake is representative of one of three phases (see subsequent figure): linear or integrative (i.e., no significant losses of accumulated residues, equilibrium not approached during the exposure), curvilinear (partly integrative, equilibrium approached) and equilibrium (the amount of chemical taken up is exactly equal to the amount of chemical lost, per unit time)  


·        Ambient exposure conditions affect sampling as described in the "SPMD sampling" subsection


·        Exposure duration is a major factor in the total amount of chemical accumulated


·        Water or air concentration of target compound also affects the amount of chemical accumulated but not its uptake rate (i.e., the daily volume of an medium cleared of chemical)

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Examples of Contaminants That are Significantly Concentrated in Triolein-Containing SPMDs (not all-inclusive)

1.        Polychlorinated dioxins and furans

2.        Polycyclic aromatic hydrocarbons (PAHs)

3.        Polychlorinated biphenyls (PCBs)

4.        Organochlorine insecticides

5.        Pyrethroid insecticides

6.        Several herbicides and many industrial chemicals

7.        Alkylated selenides

8.        Tributyl tin

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Mass of an Analyte Sequestered by an SPMD Depends on

1.        SPMD sampling rate; higher temperatures (water) and flow rates/turbulence generally provide higher SPMD sampling rates 

2.        Water or air concentration during interval of exposure

3.        Exposure duration

4.        Level of fouling or coating of the exterior membrane surface

Note: Assuming background concentrations are very low in SPMDs, the investigator can increase the number of SPMDs or exposure length (caveat:  during long exposures fouling often diminishes uptake rates) to increase the mass of sequestered analytes.

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Size and Number of SPMDs Needed for a Project Depend on

1.        Detection and quantitation limits desired by the investigator

2.        In situ SPMD sampling rate for the chosen analytes

3.        Average water concentration of the analytes during the exposure interval

4.        Sensitivity and selectivity of the chosen analytical method or bioassay

5.        Single or multiple points in time (time resolution desired)

6.        Replication or statistical requirements; the coefficient of variation (C.V.) of contiguous replicate SPMDs is typically < 20%

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General Comments on Deployment Methods


·        Typically, SPMDs are stored and shipped in clean gas-tight metal cans of various sizes


·        Metal containment structures (storage cans and deployment devices) must be free of cutting oils or other potential interferences


·        Minimize use of plastic components, except Teflon and some types of PVC, due to the possible presence of leachable organic residues


·        The structural design of the deployment device should minimize abrasion of the membrane even in turbulent environments while baffling the very high flow/turbulence of some media


·        Current velocity/turbulence is also a concern in terms of tethering, especially during floods


·        If a loop design (SPMD) is used, the two sides should not make contact


·        If water turbidity is low, then a shading structure may be required for analytes such as PAHs that undergo photolysis (caution: estimated photolysis half-lives of PAHs in direct sunlight range from 0.1 h to 5 h)


·        For sampling PAHs from air, the deployment structure must reduce ambient sunlight levels to near zero


·        Unless permeability/performance reference compounds (PRCs) are used (see description of PRCs in subsequent section), the flow and temperature regime of exposure sites should be similar to facilitate inter-site comparisons


·        Because vandalism is always a potential problem in the field, the deployment structure should be amenable to hiding


·        Deployment structures are commercially available (see "example of deployment structure")

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Examples of Deployment Apparatus

A commercially available stainless steel deployment apparatus, which has a capacity for 5 standard SPMDs.  Each SPMD is placed on a separate rack and the five racks are held in place by a threaded center pin as shown in the picture.  Also pictured are gas tight steel cans used for transport and storage.


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Recovery and Storage of SPMDs

1.        As soon as SPMDs are recovered from the environment, they should be sealed in the original can and placed on ice in a cooler for shipping (overnight shipping is recommended)

2.        Some loss of SPMD-sequestered analytes with high Henry's constants (> 10-3 atm-m3/mole) and low KOW values (£ 1 x 103) is possible but should be minimal when compared to excised water samples

3.        SPMDs should be stored in the sealed cans (shipping containers) in a freezer at -20 o C until analysis

4.        To ensure the validity of this storage method, a relatively high fugacity (escaping tendency) compound (2,4,5-trichlorophenol) was spiked into several SPMDs and the devices were left open in a freezer; no losses were measurable after 6 months

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Preparation for Analysis

1.        To permit dialytic recovery of residues in the SPMD membrane and lipid, the periphyton, mineral precipitates, and sediments/soot (air) must be removed from the exterior membrane surface (see Huckins et al., [1996] for cleaning procedure)

2.        Membrane cleaning is not necessary if only the lipid is analyzed or assayed

3.        After cleaning, check for holes in the membrane to ensure the sampler’s integrity

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Comments on Analyte Enrichment Required For Analysis or Assay of Residues in SPMDs

1.        Analyte recovery and enrichment procedures for SPMDs generally require less effort than those for tissue and sediment matrices

2.        Similar to all other environmental matrices, recovered analytes that are members of complex mixtures typically require class fractionation (not necessary for bioassay) after cleanup of any SPMD-related lipids, polyethylene waxes, and other interferences

3.        SPMD extracts are readily amenable to a classic TIE approach to elucidate the identity of toxicants

 Petty et al. (2000) have discussed analytical methods required for SPMD analysis. The following figures illustrate general procedures used for the recovery/cleanup of target analytes or unknowns from SPMDs. 

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 For more details on sample processing and enrichment, see References.

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Sampling Rate Determination

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Selected SPMD Aqueous Sampling Rates (RS, Standard 1-g Triolein SPMD) for Nonpolar Organic Contaminants1





































2, 2’,5, 5’-Tetrachlorobiphenyl



1 Flow-through (< 1 cm/sec velocity) constant concentration (100 ng/L) exposures at 18 °C; biofouling was minimized
2 Value based on 4-d exposure; uptake not linear/integrative after 6 days

Note:   for a much more extensive listing of sampling rates see Huckins et al., Guide for the use of SPMDs as samplers of waterborne hydrophobic organic contaminants, API no. 4690, In press.

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Requirements for Estimating Ambient Contaminant Concentrations from SPMD Concentrations

1.        Laboratory determination of SPMD exchange rates (uptake and elimination) and KSPMDs for analytes of interest at multiple temperatures

2.        Analyte concentrations in whole SPMDs through time is recommended when feasible

3.        Average temperature during the exposure

4.        Flow velocity

5.        Estimates of possible reduction in sampling rate due to fouling

Note: Use of PRCs should negate the need for requirements 4 and 5.

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Huckins et al. (1993) described the basic theory related to the uptake and dissipation of contaminants from SPMDs and development several mathematical models for estimating water concentrations from analyte concentrations in SPMDs.  SPMD sampling rates were demonstrated to be independent of water concentration, while the amount of accumulated residues is proportional to the concentrations of dissolved chemicals.

Modeling Assumptions and Overall Exponential Model

Using a lipid-equivalent approach to model the contribution of the membrane, and making no initial assumptions about the rate-limiting step in the overall SPMD uptake process, analyte concentrations in water and air can be computed from the following relationship.

CW, A = CSPMD / KSPMD (1-exp [-ket])                                        Equation 1

Where CW, A is the analyte concentration in water or air, CSPMD is the analyte concentration in the SPMD, t is time, KSPMD is the equilibrium SPMD-water or -air partition coefficient, and ke is the first-order loss rate constant.  This model fits the overall uptake curve (see figure illustrating the three phases of SPMD uptake), but is primarily used for compounds that reach the curvilinear uptake phase during an exposure. Based on equation 1, it appears the rate that chemicals are accumulated by an SPMD is dependent on the loss rate (ke) of residues from the device, which has units of t-1.  Actually, ke is dependent on both the uptake rate constant and the KSPMD as shown by

ke = ku / KSPMD                                                          Equation 2

where ku is the linear uptake rate constant given in L/d·g, and this case, KSPMD has units of L/g.  Isotropic exchange kinetics (IEK) are implicit in both equations 1 and 2, which means that the overall process of residue uptake and elimination obeys the same rate law, and that measured in situ PRC kes values (along with KSPMD values) can be used to predict in situ kes, and thus kus of the analytes of interest.  However, use of equation 1 without curve fitting software (requires multiple points in time) is dependent on knowledge of CSPMD, KSPMD, and ke or ku.

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Estimation of KSPMD

Because the SPMD consists of two phases (i.e., lipid and membrane) that accumulate residues, the KSPMD is given by

KSPMD = (KMW VM + KLW VL) / VSPMD = KLW (VL + KML VM) / VSPMD                        Equation 3

Where Kmw is the equilibrium membrane-water partition coefficient, Vm is the volume of the membrane, KML is the equilibrium membrane-lipid partition coefficient, VL is the volume of the SPMD lipid, and VSPMD is the total volume of the SPMD.  Unfortunately, only a few values of KSPMD are available for air.  By assuming that KSPMD remains constant within the typical range of environmental exposure temperatures, Equation 3 and two regression models developed by Hofmans (1998) can be used to compute KSPMD.  These regression models are as follows:

log KLW = -0.1257 (log KOW)² + 1.9405 (log KOW)-1.46                        Equation 4


log KMW = -0.0956 (log KOW)² + 1.7643 (log KOW) –1.9                        Equation 5

Note that the accuracy of the equations above diminishes at log Kows > 6.0 and that SPMD aqueous partition coefficients can be converted into SPMD air partition coefficients by dividing each value by the corresponding non-dimensional Henrys constant (H'), which are generally available in the literature.

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Linear Uptake Model (Integrative Sampling Phase)

For high KOW compounds (i.e., log KOW > 5.0), uptake is generally linear during non-turbulent, moderate temperature exposures (note that biofouling impedance in some aquatic environments may cause a deviation from linearity during longer exposures).  When uptake is linear, CW, A can be derived from CSPMD by the following relationship

CW, A = CSPMD MSPMD / Rs t = CSPMD / ku                                    Equation 6

where MSPMD is the mass of the SPMD in g and Rs (L/d) is operationally defined as the sampling rate of a "standard" 1-g triolein SPMD.  The advantage of this type of sampling (integrative) is that residues are accumulated through time without any significant losses (permits the maximum sequestration of target compound mass), and estimates of analyte levels from Equation 6 represent true TWA concentrations.

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Equilibrium Model

For compounds with moderate to low KOW s (i.e., log KOW < 5.0), and when exposure conditions are relatively turbulent and warm, residue concentrations may closely approach (i.e., ³ 90 %) or reach equilibrium concentrations in SPMDs.  In this case, the following model is applicable to the estimation of ambient environmental concentrations.

CW,A  = CSPMD-e /KSPMD                                                Equation 7

where CSPMD-e is the equilibrium concentration of an analyte in the SPMD.

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Estimation of Times for the Phases of SPMD Uptake

In the absence of PRC data, selection of the appropriate model for estimation of ambient concentrations from SPMD concentrations is problematic.  However, the following equations can be used to provide some guidance.

t50 = t1/2 = -ln 0.5 KSPMD VSPMD / Rs = -ln 0.5 KSPMD / dSPMD ku        Equation 8  

t90 = -ln 0.1 KSPMD VSPMD / Rs = -ln 0.1 KSPMD / dSPMD ku                Equation 9  

t1/2 » -ln 0.5 KOW VSPMD / Rs » -ln 0.5 KOW / do ku                        Equation 10

where t50 is the time required to accumulate 50 % of the equilibrium concentration, and is mathematically identical to t1/2 (time required to lose 50 % of the initial residue concentration), t90 is the time required to reach 90 % of the equilibrium concentration, and dSPMD and do are the density of the whole SPMD (» 0.91 g/cm3) and octanol.  Note that the first 50 % of the total residue accumulated essentially represents the linear region of uptake, > 50 % to 90 % represents the curvilinear region, and > 90 % represents the steady state or equilibrium region.  If the KSPMD and KOW are written as unitless coefficients, the density corrections are unnecessary.  KOWs are widely available and a considerable number of SPMD Rs and/or ku values are now available in the "Guide For The Use Of Semipermeable Membrane Devices (SPMDs) As Samplers Of Waterborne Hydrophobic Organic Contaminants" by Huckins et al. (2001).  See the following subsection on PRCs for a discussion on how the amount of PRC loss during an exposure can be a guide for the choice of the appropriate model for water or air concentration estimates.

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Permeability/Performance Reference Compounds

As previously indicated, PRCs are (analytically) non-interfering compounds with moderate to relatively high fugacity from SPMDs, which are added to the SPMD lipid prior to deployment.  We use "permeability" to refer to compounds under membrane control and "performance" to refer to compounds under external boundary layer control.  The use of PRCs can be viewed as an in situ calibration/recalibration approach, where the rate of PRC loss during an exposure is related to the target compound uptake.  This is accomplished by measuring PRC loss rates (kes) during calibration studies and field exposures.  Using these values, an exposure adjustment factor (EAF) can be derived.  The EAF, along with kus and kes from calibration studies can then be used to predict in situ SPMD kus of environmental contaminants.  A fundamental assumption of the PRC approach is that the EAFs of PRCs with log KOWs generally < 5.0, can be used to predict the EAFs of chemicals with much higher log Kows.  Based on a recent study by Huckins et al. (2002), this assumption appears valid and the difference between measured concentrations of an analyte and the PRC derived estimates should be within two fold.

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When temporal losses of PRCs are measured (n > 3), regression analysis can be used to determine PRC ke values.

CSPMD = CSPMD-0 exp (-ke-PRC t)                                               Equation 11

where CSPMD-0  is the initial PRC concentration.  Generally, a field blank containing a PRC is used to determine CSPMD-0.  If PRC levels in SPMDs are measured only at the beginning and the end of a field exposure, Equation 11 can be solved to permit a two-point derivation of ke-PRC s (assuming first-order kinetics) as follows:

ke-PRC = ln (CSPMD-0 / CSPMD) / t                                              Equation 12

and the EAF is derived by

EAF º ke-PRC-f / ke-PRC-cal » (ku-f / KSPMD -f) / (ku-cal / KSPMD-cal)                          Equation 13

where “-cal” and “-f” refer to values measured during calibration studies and in the field.  The ku-f (i.e., an estimate of the actual in situ sampling rate of the target compound during a field exposure) can be derived by

ku-f » KSPMD-f EAF (ku-cal / KSPMD -cal)                               Equation 14

Selection of compounds to serve as PRCs is limited by the need to have measurable losses of PRC residues during an exposure and the ability to differentiate PRC residues from other quality control standards, target compounds and unknowns of potential interest to an investigator.  Depending on the Kows of target analytes, candidate PRCs may have to include compounds that are representative of both membrane and diffusion control.  Also, some information on general environmental conditions (e.g., flow rate and temperature) at sample sites and the duration of planned exposures are advisable to help ensure that an acceptable range of PRC loss occurs during exposures.  For example, PRC losses are enhanced under exposure conditions of warm turbulent waters.  To prevent the complete loss of PRCs under this scenario, the use of compounds with moderately high log Kows (i.e., range of 4.5 to 5.3) may be necessary and/or exposure length may have to be shortened.  In addition, larger quantities of these PRCs and those with low log KOWs (i.e., < 3.5) may have to be spiked into SPMDs.  These precautions are necessary to ensure that changes in PRC residue concentrations can be statistically delineated from the C.V.s (%) for SPMD sample analyses.  Even when PRC loss or retention is too great to use for the derivation of EAFs, information on the kinetic phase of analyte uptake (see figure on the three phases of SPMD uptake) at the termination of the exposure is still gained (Booij, 2000).  For example, if a PRC with a log Kow of < 4.5 is completely lost during an exposure, then all analytes with log KOWs of < 4.5 should have attained equilibrium (i.e., KSPMD-e).  On the other hand, if no loss of a PRC with a log Kow of > 5.0 is observed then linear uptake can be assumed for all analytes with log Kow of > 5.0.

The use of certain perdeuterated (all hydrogen atoms replaced with deuterium atoms) compounds appears to offer considerable promise as PRCs.  This type of labeled compound is commercially available at relatively low cost (when compared to similar 13C-labled compounds), has physicochemical properties (excluding molecular weight) similar to their native analogues, is not found at significant levels in the aquatic environment, and generally can be separated from their native analogues by high-resolution gas chromatography.  Based on our experience, the following perdeuterated PRCs are recommended: naphthalene, acenaphthene and fluorene for membrane controlled analytes and phenanthrene, anthracene and pyrene for diffusion layer controlled analytes.  Note that if PAHs are used for PRCs or are target analytes, exposure of SPMDs to sunlight must be avoided because they readily photodegrade.  When analyses are performed by mass spectrometry, then it is feasible to use 13C -PRCs if appropriate standards can be found at a cost within budget of the project.  Obviously, non-labeled compounds, such as 2, 2'-dichlorobiphenyl and 2, 4, 5-trichlorobiphenyl (not present in commercial PCB mixtures), can be used for PRCs as long as they are not present in the environment sampled and are not used as analytical quality control standards.

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SPMD-Biota Comparisons

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These two plots show the similarity of the uptake of a wide range of contaminants by SPMDs and the gills of fish.  In both cases, uptake rates peak between log KOWs of 5.0 and 6.5 and the overall shape of the curve appears to be parabolic.

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This plot shows that for high KOW compounds such as PCBs, uptake is similar and linear (integrative) for both SPMDs and fish. However, standard-SPMD uptake rates of total PCB congeners were about 1.8 fold higher than uptake rates of the test fish and the capacity (i.e., equilibrium mass of analyte accumulated divided by mass of the sampling matrix) of SPMDs is greater than fish.

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Numerous chromatograms of SPMD extracts, such as those above, show proportionally greater quantities of nonpolar low molecular weight or low KOW chemicals sequestered in SPMDs than in biota.

Because of the seeming bias (relative to most biomonitoring organisms) of SPMDs to accumulate larger amounts of low KOW compounds (log KOW < 4.5) relative to high KOW compounds (log KOWs > 6.0), the following question arises:  are SPMD sampling rates for hydrophobic chemicals of low molecular weight greater than those for hydrophobic chemicals of moderate-to-high molecular weight?  Before answering this question, we should review the meaning of “sampling rate”.  SPMD sampling rates, Rs and ku, are given in Ld-1 (Rs) or Ld-1g-1, which are first-order rate constants independent of water concentration.  These rate constants represent a characteristic volume of water that is daily cleared of chemical by an SPMD (lipid plus membrane).

SPMD Rs and ku values have been shown to rise with KOW or molecular size until boundary layer control is evoked (log KOW » 4.5). Then, falling diffusion rates with increased MW and potential solubility and sorption limitations, significantly reduce RSs or kus (appears to be evident at log KOW » 6.0).  In general, sampling rates for low KOW compounds are still less than those for moderate to high KOW compounds.  However, this observation does not mean that greater amounts of low KOW chemicals will not be sequestered when water concentrations are much higher.  For example, environmental water concentrations of low KOW compounds may often be 1000-fold greater than high KOW compounds, whereas the SPMD ku values for the same compounds differ by only about 30-fold.  The amount of analyte recovered in an SPMD (Ma) is given by

Ma = ku MSPMD t Cw                                                                 Equation 15

Where Ma is the mass of analyte sequestered.  Therefore, it is not surprising that chromatograms of SPMDs generally have higher amounts of lower KOW or more water-soluble compounds.

Also, biota more readily metabolize or depurate most low KOW compounds.  Finally, if one allows sufficient exposure time for SPMDs to approach equilibrium for all accumulated compounds, differences between the chromatographic profiles of low KOW and high KOW chemicals (sampled earlier) should be reduced.

Because many types of environmental stressors can affect the health, and thus the dietary and respiratory uptake of chemicals by transplanted biomonitoring organisms, a disparity may exist between the number and relative amounts of analyte residues detected in SPMDs and the tissues of test animals.  The following figure illustrates this difficulty.  

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This side-by-side exposure of SPMDs and clams shows that the number and amounts of chemicals accumulated by SPMDs and biomonitoring organisms are not always comparable.  In the case shown above, clams accumulated only a small fraction of the PAHs residues concentrated by the SPMDs.  The investigators (Moring and Rose, 1997) suggested that the clams were not filtering water (feeding) during most of the exposure, because of stress.

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Use Of SPMDs In Conjunction With Biomarkers/Bioindicators

Use of certain in vitro biomarker/bioindicator assay to assess ecosystem health is sometimes problematic, because of the difficulty of obtaining samples suitable for testing.  Most hydrophobic organic contaminants are present in environmental water only at trace levels (i.e., < 1 µg/L).  However, the sometimes-slow process of bioconcentration (uptake from water, respiratory and dermal absorption [e.g., non-scaled fish])/bioaccumulation (dietary, respiratory, and dermal absorptive uptake) can lead to elevated concentrations of contaminants in aquatic organism tissues, which can result in a variety of adverse effects.  In some cases, samples and procedures used for cellular-based assays may not account for the potential effects of bioconcentration.  Also, some bioindicator tests have relatively low sensitivities for many pollutants.  Thus, direct testing of environmental waters with these assays may lead to false-negative errors in assessing the potential risk of waterborne residues to aquatic life.  To avoid this type of error and expand the use of these tests for ranking toxicity potential, a preconcentration method is often needed that mimics the biouptake process.

Complex mixtures of chemicals are mimetically sequestered by SPMDs and are often amenable to examination by a variety of bioassays.  These assays include both biomarker/bioindicator tests and immunoassays.  Assays that have been used to assess SPMD extracts or diluents include the following: Microtox®, Mutatox®, mixed-function oxygenase induction-ethoxyresorufin-o-deethylase (EROD) activity, sister chromatid exchange, vitellogenin induction via interperitoneal injection of test species, enzyme-linked immunosorbent assay, and Ames mutagenicity test (note that this list is not all inclusive).  The marriage of SPMDs and compatible biomarker/bioindicator tests offers many avenues of investigation, all potentially providing information concerning the relative toxicological significance of chemicals present in the environmental matrices sampled.

However, two issues should be considered to ensure that the results of SPMD-biomarker/bioindicator tests provide toxicologically relevant information.  First, the level of residue preconcentration by the SPMD should be less or fall within a range of estimated bioconcentration/bioaccumulation levels in feral organism tissues.  Secondly, the levels of oleic acid and elemental sulfur in SPMD extracts should be kept at a minimum.  Fatty acids are known to be cytotoxic and sulfur may be reduced to toxic sulfides (Brouwer and Murphy, 1995).  The toxicity of fatty acids such as oleic acid has been attributed to their membrane disturbing properties, which include disruption of the calcium pump by the formation of metal salts (Sabaliunas et al., 2001).  Oleic acid is the only fatty acid present as an impurity in the 95% triolein commonly used in SPMDs.  During environmental exposures, a significant portion of this triolein impurity diffuses to the exterior surface of an SPMD, where dissipation and/or degradation occur.  Unfortunately no or little attenuation occurs in the oleic acid levels in laboratory SPMD-field blanks, -fabrication blanks and -process blanks.  Thus, the potential for significant differences in biomarker/bioindicator test responses exist among reference site (i.e., no toxicant present) exposed SPMDs and associated field and laboratory QC SPMD samples.  Fortunately, only extremely low levels of oleic acid are present in some commercially available triolein (e.g., Sigma, 99 %, and Nu-Chek Prep., Inc., 99 %). If cost limitations require the use of triolein with oleic acid, a simple silica gel cleanup step can be used to separate oleic acid from target pollutants in SPMD extracts (contact Jon Lebo, CERC, USGS, Columbia, MO, USA, ph: 573-876-1837) and may be scaled up to purify triolein prior to use in SPMDs.  Also, it is noteworthy that fatty acids are commonly present in extracts of fish and other aquatic organisms. In the case of sulfur, many types of sediment contain relatively large amounts of elemental sulfur and elemental sulfur is readily accumulated by SPMDs.  To ensure that the results of biomarker/bioindicator tests reflect the effects of target compounds, elemental sulfur can be removed from extracts by treatment with size exclusion chromatography or with shiny copper (contact Jon Lebo, CERC, USGS, Columbia, MO, USA, ph: 573-876-1837).

The following figure and table illustrate the application of SPMDs as mimetic sample concentration/collection devices:

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Use of Microtox® and Mutatox® to Determine the Toxicity of SPMD Concentratesa.  Microtox values are 5-minute EC-50s with 95% confidence intervals (in parentheses); Mutatox values show positive response as "+" and negative response as "-" (courtesy of Tom Johnson). 


Sample Type

Microtox Toxicity (EC-50)a

Mutatox Genotoxicity (+/-)




Winter Quarters Bayb



McMurdo Soundb



Flat Branchc



Quality Control



Procedural Blanke



SPMD Controlg



Microtox phenol reference toxicant (µg/mL H2O)



Mutatox benzo-a-pyrene reference toxicant

(1.0 µg/vial)




a Assays were conducted on lipid diluent or dialysates and EC-50 values represent mg SPMD lipid/mL carrier solvent

b SPMDs were exposed to Antarctica sediments in microcosms
c SPMDs were exposed to a small urban stream
d None analyzed
e Solvents/reagents used in tests
f None detected
g Freshly prepared SPMD; carried through Microtox and Mutatox test

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The figure shown below demonstrates that PCB vapors (top chromatogram) are readily sampled by SPMDs.  In fact, the potential utility of SPMDs for atmospheric sampling (e.g., Petty et al., 1993 and Ockenden et al., 1998) is as great as that of water.  In general, the linear uptake rates (i.e., volumetric Rss and kus) are about a thousand times higher in air than in water.  The much larger volumes of air sampled, relative to water, are primarily due to the approximately 10³ higher diffusion coefficients of vapors relative to aqueous solutes.  However, when the difference in the density of water and air is considered, the adjusted sampling rates for the two phases are of similar magnitude.  Note that we evoke differences in media density solely for comparative purposes, as density corrections are not performed in interphase mass transfer models.

There is little calibration data (standard SPMD sampling rates) in the literature other than those of Ockenden et al., 1998.  The results of the Ockenden et al., 1998 and unpublished work by CERC scientist, indicated that for a wide range of PCB KOWs  (exposure condition: low air velocity at 4 °C to 26 °C), the Rss for standard SPMDs varied from about 0.5 m³/d to 20 m³/d.  Currently, additional calibration studies are underway at CERC.  Similar to water, the rate-limiting step in the uptake of hydrophobic chemicals may be the external boundary layer, because the "effective" mass-transfer resistance in the membrane is less in the air than in water, as values of a chemical's KOA are generally much higher than it's KOW.

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Sediment Pore Water

The table below shows that SPMDs sample sediment associated contaminant residues.  Exposure conditions in the bed sediments of many aquatic environments are less variable than in the water column.  Therefore, the sometimes-large differences in the sampling rate of a target compound in the water column at different sites (largely due to changes in hydrodynamics) should not materialize in bed sediments.  On the other hand, the heterogeneity of the amount and quality of sediment organic carbon and textural differences at sample sites likely plays a major role in the amount of chemical accumulated in SPMDs.

When using SPMDs to sample the water column of aquatic systems, chemical concentrations in the outer portion of the device’s aqueous boundary layer reflect residue levels in the bulk medium, because of rapid mixing-exchange of residues from the water column.  In the case of SPMDs placed in sediments, resupply of hydrophobic residues extracted from the SPMD-associated aqueous boundary layer and/or contiguous pore water may be limited by the rate of residue desorption from sediment particles and/or slow diffusion in stagnant pore water and in sediment organic matter. Comparison of chemical concentration factors (CFs) in the triolein (i.e., residue concentration in SPMD lipid divided by the residue concentration in the water column just above the sediment) of SPMDs covered by sediments to those measured in SPMDs placed in slowly flowing water free of sediments (similar temperatures) showed that the CFs in sediment-exposed SPMDs were about half of those observed for SPMDs exposed in water alone (see table below for CFs of selected compounds in SPMDs in sediment).  Note that this potential discrepancy should disappear if actual pore-water concentrations were known.  This data suggests that the concentration of 2, 2', 5, 5'-TCB in porewater contiguous to the SPMD was significantly reduced relative to the water column.  In general, the concentrations of chemicals in relatively undisturbed pore water of benthic sediments are expected to be greater than the concentrations of the same compounds in the water column.  Until SPMDs reach equilibrium with contiguous sediments, estimation of the "undisturbed" porewater concentrations of high KOW compounds is not possible.  However, use of solid phase microextraction (SPMEs) fibers with very thin films (e.g., 7 µm film thickness of polydimethysiloxane [PDMS]) of sorbent may permit estimation of non-disturbed pore water concentrations.  These thin-film SPMEs have high surface area for solute exchange and low-sorbent volume, which reduce the time required toe reach equilibrium with sediment-associated residues.  Once equilibrium is achieved, computation of undisturbed pore water concentrations is straightforward.

Although SPMDs generally will not reach equilibrium with pore water, they provide valuable information on the amount of bioconcentratable residues available during a window of time.  Perhaps the best approach for the characterization of contaminated sediments is a two-part assessment, where SPMDs are used in situ to measure the flux of bioavailable chemicals (results relate to the toxicity of sediments to laboratory test organisms), and to detect and collect relatively large amounts of trace-level bioavailable contaminants (may be used for bioassay or toxicity identification evaluation procedures).

                                Exposure of SPMDs1 to Contaminated Sediment2


Concentration Factor (CF) in Triolein


1 day

7 days

14 days

28 days











2, 2', 5, 5'-TCB 3





1 SPMDs (n = 3/test chemical) contained 0.1 g triolein and were covered by sediment
2 300-g sediment (% organic = 1%) and 950 mL CERC well water
3 Tetrachlorobiphenyl

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The table below shows that SPMDs and aquatic organisms often have similar uptakes rates (kus) for hydrophobic organic chemicals, but the clearance rates (kes and t1/2s) of accumulated residues from organisms are generally much faster than SPMDs (see table below and the two related figures that follow).  Remember that integrative sampling requires that sampling is additive through time, i.e., once residues are taken up they are not lost when environmental concentrations vary. The observed difference in the residence times of residues in the two matrices is due to several factors.  Only relatively slow, surface area dependent, passive diffusive exchange processes govern the rates of residue losses from SPMDs, whereas residue-loss rates from aquatic organisms are governed by much more complex processes.  Active respiration, feeding, waste elimination, metabolism, reproduction (females) and passive diffusion combine to control the depuration of accumulated contaminant residues from organisms.  Obviously, another major difference between SPMDs and nearly all aquatic organisms is that SPMDs have a much larger lipid pool than organisms of equivalent mass and, unlike organism proteins, the LDPE membrane significantly adds to the capacity of the sampler.

Comparison of SPMD and Organism Uptake (ku) And Clearance (ke) Rate

Constants for Selected Chemicals






SPMD (membrane + lipid)


 ku(Ld-1g-1)      ke(t-1)          t1/2*

  Organism (whole body)  

 ku(Ld-1g-1)        ke(t-1)             t1/2




32 h







(Daphnia pulex)




0.4 h




3.5 h




240 h







500 h



(Daphnia pulex)

1.3 h

2, 2', 5-trichlorobiphenyld










(Brown Trout)








346 h

2, 2', 5, 5'-tetrachlorobiphenyle



1,730 h





(Brown Trout)



(Zebra Fish)





433 h

a SPMD: This work; MacKay et al., 1992, organism not specified
b SPMD: This work  
c SPMD: This work; Biota, MacKay et al., 1992
d SPMD & Trout: Meadows, CERC, 1996;Goldfish, MacKay, et al., 1992
e SPMD & Trout: Meadows, CERC, 1996; Zebra Fish, MacKay, et al., 1992
* First-order half-lives

The much longer half-lives and kes shown in the table above indicate that SPMDs are better suited for integrative sampling of contaminant residues.  The passive integrative sampling of high KOW compounds by SPMDs enables detection of episodic releases and provides more accurate time weighted average estimates of water concentrations.  On the other hand, biomonitoring organisms are more likely to reach steady state concentrations, may sample ionic compounds (unlike SPMDs), and the analysis of feral organisms is essential for determining residue concentrations in edible tissues.  However, residue concentrations in biomonitoring organism tissues often may not be proportional to ambient environmental concentrations of contaminants, which suggest that tissues levels cannot reliably be used to derive water concentrations and thus differences in bioavailable pollutant levels at study sites may often be difficult to discern.

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Use of lipid normalization in environmental sample analysis must be approached with caution, especially when comparing pollutant concentrations in lipid containing SPMDs and biomonitoring organism tissues.  The lipid content (%) of aquatic organisms used in environmental studies varies considerably but generally falls in the range of 0.2 to 10 %, whereas SPMDs are » 20 % lipid.  Also, the SPMD membrane has a much higher capacity for hydrophobic chemicals than non-lipoidal tissues of organisms.  The previous table clearly shows that times to equilibrium for SPMDs and biomonitoring organisms exposed to hydrophobic contaminants are usually quite different.  A key assumption of lipid normalization theory is that all sample matrices are at equilibrium with ambient contaminant residues.  The following illustration shows how errors are generated, when lipid normalization is applied to a sample that has failed to reach equilibrium with the ambient environment.

Comparisons of SPMDs and Biota (Kinetics and Steady State)

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Requirements for Lipid Normalization:


·        Samples must be at steady state or equilibrium with ambient environment (seldom the case with SPMDs)


·        Solvents and methods used for sample extraction must be similar to ensure recovery of the same types and amounts of sample lipid

Assumptions of Lipid Normalization:


·        Differences in lipid composition among individuals and species have little or no effect on the magnitude of equilibrium lipid/water partition coefficient


·        Variance is decreased relative to non-normalized data


Lipid normalization is often inappropriate, especially in the case of SPMD-biota comparisons, largely because SPMDs seldom reach steady-state concentrations.  

For Further Information

Although this tutorial covers many of the key elements of SPMD technology, it is not comprehensive and certain aspects will be updated as ongoing research is completed.  The annual or semiannual International SPMD Workshops are excellent sources of information on current developments in SPMD technology.  (Contact Environmental Sampling Technologies, 1717 Commercial Drive, St. Joseph, MO 64503.)

If the reader has additional questions concerning SPMDs, contact Dave Alvarez: (573) 441-2970; e-mail:

We have also prepared answers to 20 frequently asked questions (FAQs) related to SPMD technology.

See References for peer-reviewed journal articles about various aspects of Semipermeable Membrane Devices and their applications and for articles referred to in the tutorial.  Also, the reader should be aware of an extensive forthcoming report for the American Petroleum Institute (API):   "Guide for the Use of SPMDs as Samplers of Waterborne Hydrophobic Organic Contaminants," API no. 4690, In Press.  This document expands the discussions herein, provides an extensive set of calibration data for standard SPMDs, and covers topics not discussed in this tutorial.

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