Sediment Conventionals

Grain size, organics and stable isotopes

Field, laboratory and QA/QC protocols for measuring some of the recommended sediment conventionals are described in detail in PSEP 1986 (Recommended Protocols for Measuring Conventional Sediment Variables in Puget Sound:

Typically, a dedicated grab sample is obtained for sediment conventionals. The top two to three centimeters of sediment will be collected with a stainless steel spoon from the center of the grab for TOC, grain size, and chemistry analyses. The sediment will be put in a stainless steel bucket and covered with a lid. On subsequent grabs, the top two to three centimeters of sediment away from the sides of the grab will be collected and added to the bucket. Grabs will be taken until enough sediment is collected to fill all necessary sample containers for the station.  The composited sediment in the bucket will be homogenized by stirring with a stainless-steel spoon or paint mixer until a uniform texture and color are achieved. After the sample jars are filled. Leftover sediment will be returned to the water column at or near the sites where collected. At 5% of the stations sampled, double the amount of sediment will be collected and homogenized, with a second set of sample containers for chemistry, TOC, and grain size analyses. The second set will be assigned a different sample identification number and submitted to the laboratories as a blind field replicate. 

A portion of each sample will be jarred and retained as grain size and TOC/chemistry archive samples. They will be kept for one year, in case re-extraction or retrospective analysis is required. Sediment grain size samples will be held at 4 °C. Chemistry and TOC samples will be frozen at –18 °C (0 °F). 

1. Grain Size analyses

Particle size determinations can either include or exclude organic material. If organic material

is removed prior to analysis, the “true” (i.e., primarily inorganic) particle size distribution is

determined. If organic material is included in the analysis, the “apparent” (i.e., organic plus

inorganic) particle size distribution is determined. Because true and apparent distributions may

differ, detailed comparisons between samples analyzed by these different methods are

questionable. Therefore it is recommended that all samples be analysed without organics. This can be accomplished by picking out larger, obvious infauna. 

Samples can be collected in glass or plastic containers. A minimum sample size of 100-150 g is

recommended.  Samples should be stored at 4° C, and can be held for up to 6 mo before analysis. Samples must not be frozen or dried prior to analysis, as either process may change the particle size distribution. Detailed laboratory methodology and QA/QC is included in PSEP (1986).

Particle Size Distribution — These analyses will be carried out using a method described in Walton (1978), to determine the proportions of material present in each of the following particle size categories: Gravel (>2.00 mm)

Gravel (>2.00 mm) Coarse Silt (<0.0625 – 0.0312 mm) 
Very Coarse Sand (<2.00 – 1.00 mm) Medium Silt (<0.0312 – 0.0156 mm) 
Coarse Sand (<1.00 – 0.500 mm) Fine Silt (<0.0156 – 0.0078 mm) 
Medium Sand (<0.500 – 0.250 mm) Very Fine Silt (<0.0078 – 0.0039 mm) 
Fine Sand (<0.250 – 0.125 mm) Clay (<0.0039 mm) 
Very Fine Sand (<0.125 – 0.0625 mm) 


2. Sediment pH

This analysis will be carried out according to APHA Method 4500-H ―pH Value (1998), using a pH electrode. 

3. Total sediment dry weight (inverse of moisture content)

Total dry weight of sediments is determined by removing sea salt by washing and centrifugation with distilled de-ionized water.   After rinsing, the samples were freeze dried and gently disaggregated in an agate mortar and pestle and transferred to a clean plastic vial.  The dried sample is used for the total carbon, total nitrogen, carbonate carbon, biogenic silica, d13Corganic and d15Ntotal analysis.

4. Total carbon, organic carbon, total nitrogen and carbonate analysis, including biogenic silica and opal

Samples can be collected in glass or plastic containers. A minimum sample size of 25 g is recommended.  Total organic carbon should be reported as a percentage of the dry weight of the unacidified sample to the nearest 0.1 unit.  A complete description of analytical methods is given in PSEP (1986) and Calvert et al. (1995).  

Wet sediment samples are dried in a freeze dryer, then finely ground with a Hertzog grinding mill.  About 15-20mg of ground sample is weighed into tin cup.  The tin cup is placed into an autosampler on the Elemental Analyzer for analysis as described below.

Total carbon, total nitrogen and carbonate analyses are typically performed using combustion/gas chromatography such as a Carlo Erba CHN analyzer (Verardo et al. 1990; King et al. 1998).  If the elemental analyzer used to measure total organic carbon can also measure total nitrogen, it is recommended that the latter variable be measured simultaneously with TOC, as follows;

Samples are introduced to the CN Elemental Analyzer by an autosampler.  The sample is dropped into a quartz combustion column heated to 1000 ºC along with a stream of oxygen which produces a flash combustion of the sample, forming a mixture of CO2 and NOx. The combustion column is packed with lead chromate, copper oxide wires and aluminum oxide.  The sample gases are carried through the combustion column by helium carrier gas into a reduction column.  The reduction column is filled with metallic copper which removes the excess oxygen and the nitrogen oxides are reduced to N2.  The N2, CO2, and water from the reduction column are swept through a water trap containing phosphorus pentoxide.  The gases are separated by means of a thermally programmed GC desorption column.  The NOx and CO2 are adsorbed by the column, then desorbed sequentially by heating the column to programmed temperature steps.  Standards with known amount of C and N are analyzed along with the samples to produce a standard curve which allows conversion of the instrument units to microgram of TOTAL C and N.

Carbonate or inorganic carbon is separately determined by acid evolution of CO2 and quantification using a UIC coulometer (Huffman 1977; Johnson et al. 1993; precision ± 1.6%, 1 standard deviation).  This process uses a UIC, Inc. CM5130 Acidification module to acidify samples in order to evolve forms of inorganic carbon as carbon dioxide.  This CO2 evolved is then carried through a scrubbing system by CO2-free gas stream into a Model 5014 CO2 Coulometer for detection.  The CO2 gas stream passes through the coulometer cell which is filled with monoethanolamine and a colorimetric pH indicator.  The CO2 is absorbed and reacts with the monoethanolamine to form a titratable acid.  This acid causes a color indicator in the coulometer cell to fade.  A photodetector monitors the color changes as a percent transmittance (%T).  As the %T increases, a titration current is automatically activated to electrochemically generate base at a rate proportional to the %T.  The current stops when the generated base causes the solution to return to its original color or %T.  The total cell current is integrated and displayed on a digital readout.  Reference samples of calcium carbonate are analyzed regularly to ensure proper instrument operation.   

Total Organic Carbon for the sediments, the organic carbon is achieved by taking the difference between the total carbon and inorganic carbon after the method of Calvert et al., (1995).  The filters were pre-acidified to remove inorganics prior to analysis so the analyzed carbon is already the organic carbon.  

Biogenic Silica is determined following the method and equations of (Mortlock and Froelich 1989 and see Calvert et al., 1995).  All sampling and processing equipment must be plastic, with no glass.  The method consists of extracting amorphous silica from a sediment sample with 2M Na2CO3 and then measuring the dissolved silicon concentration in the extract by molybdate-blue spectrophotometry.  This also follows procedures adapted from APHA Method 4500-Si ―Silica. 

Weigh samples into round bottom centrifuge tubes.  Weights are generally in the 20mg range.  Use weighing papers and forceps.  Add 2ml 10% H2O2 (to dissolve organics) and swirl each tube.  Wait ½ hr, add 2ml 10% HCl (to dissolve inorganics) to each.  Sonicate for ½ h.  Leave caps lightly placed on tubes.  Work in sets of 32 samples as the sonciator can only hold 32 tubes at a time. After sonicating, add 20ml DDW using the Eppendorf pipette.  Wash down sides of the tubes while dispensing.  Centrifuge for 10mins at high setting.  Decant and dry at 50c in oven overnight.  Leave caps on the tubes ajar to allow evaporation.  

Next day, add 20ml 2M Na2CO3 to each tube and put in an 85oC water bath.  1.5h after the samples have gone in the bath, vortex each tube.  1h after that, swirl each tube.  1hr after that, swirl again.  Leave in water bath for another 1.5h without disturbing the sediment.  The total time in water bath is 5 hrs. 

Arrange redox bottles in numerical order, along with standards bottles.   Dispense 10 ml DDW to each bottle using an adjustable 10ml pipette.  Extract 100 ul from each standard into the appropriate bottles before the tubes come out from the bath. Extract 100 ul from each sample tube into the corresponding redox bottles (30ml nalgene bottles) immediately coming out of bath.  Put all tubes back in bath so that extraction can be done again in case of a problem. Add 4ml of oxidant in 10 secs interval.  Make sure that it’s 20 mins between the addition of oxidizer and the 6 ml addition of reducer.  Leave caps on bottles at all times and use an adjustable pipette for dispensing. Leave samples to color in dark.  Read at 812 nm on the spectrophotometer the next day.

OXIDIZER – – mix Ammonium Molybdate (16.0 g (NH4)6Mo7O24. 4H2O) to 1 L of DDW, stored in a tightly capped poly. bottle; if white discard) and HCl  (48 ml 12N HCl + 952 ml DDW) at a ratio of 1:1 just before adding to samples (good for 6-12 hrs).

REDUCER – – mix metol-sulfite (12g Na2SO3 /L DDW + 20g Metol (p-methylamoniphenol sulfate; store in ground glass stoppered bottle in dark.  Good 1-2 months, discard if light brown color) , oxalic acid (60g COOH2.5H2O/L DDW), and sulfuric acid (300 ml conc. H2SO4 + 770 ml DDW) in a ratio of  1:1:1 ( in respective order)

5. Acid Volatile Sulphide (AVS) and Simultaneously Extractable Metals (SEM) 

Acid-volatile sulfide is defined operationally as the sulfide fraction that is evolved from sediment when treated with acid. It is a complex and variable fraction of sediment represented by a variety of reduced sulfur components, although often dominated by relatively labile Fe and Mn monosulfides. Greater AVS concentrations are associated typically with organic-rich, anoxic deposits and lower levels are found usually in oxic sediments having low organic content. Hammerschmidt and Allen (2010) inferred from that large interlaboratory variations of AVS and SEM result from differences in the method by which they are extracted from sediments in each laboratory. Such variability results most likely from either AVS oxidation during sample preparation/analysis or operational differences in extraction.  Thus the laboratory protocol must reduce oxidation potential during handling. 

Sediment samples are placed into glass jars and sealed with Teflon caps with no airspace above the sediment, to minimize AVS oxidation during transportation and storage The sample jars are then placed into an ice-filled cooler for transport to the laboratory. Sediment AVS is stable for at least 56 d when stored either refrigerated or frozen (Allen et al., 1993), but extractions should  be performed within 18 d of receipt. 

AVS can be determined using inductively coupled plasma mass spectrometry  (ICP-MS) and methods can be found Gobeil et al. (2001) or USEPA (2005). AVS and SEM extraction with 1 N HCl has been proposed, in part, to promote greater intercomparability of AVS and SEM results between laboratories (Allen et al., 1991). Sediment samples for SEM/AVS analysis are placed as collected into a cold 1 mol/L hydrochloric acid solution in a purge and trap system. The evolved hydrogen sulphide is carried into a basic zinc acetate solution by argon gas. The evolved hydrogen sulfide (H2S) gas from the sediment and acid mixture is captured and analyzed with a gas chromatograph using a photoionization detector. The AVS is thus determined colourimetrically (Allen et al. 1991). The cold hydrochloric acid extracts from the samples are then analyzed using ICP-MS for cadmium, copper, lead, nickel and zinc using EPA Method 200.8 (USEPA 1991) and using cold vapor atomic absorption spectroscopy (CVAA -USEPA Method 7471A). for mercury (Bloom and Crecelius 1983) SEM. following methods from CSR Analytical Method: Strong Acid Leachable Metals (SALM) in Soil (BC MOE, 2009;

Methods and Detection Limits for Chemistry Analyses Performed on Water and Sediment Samples

Parameter                                                        Units                DLs                  Method

Stable C/N isotopesn/a0.2 pptMass Spectrometer
Moisture Total Organic Carbon (TOC) Total Organic Nitrogen (TON) Total Volatile Solids (TVS) Particle Size Distribution Acid Volatile Sulphide (AVS) Simultaneously Extractable Metals (SEM) SEM CadmiumSEM Copper SEM Lead SEM MercurySEM Nickel SEM Zinc % wet % dry % dry % dry % dry umol/g dry μmol/g dry       0.1 0.1 0.01 0.1 0.1 0.2  0.0050 0.010 0.020 0.000050 0.050 0.0050 GravimetricCombustion Combustion GravimetricGravimetricColourimetric ICP-OES ICP-OES ICP-OES CVAFS ICP-OES ICP-OES



6. Carbon and Nitrogen stable isotope analyses

Samples can be frozen for archiving and/or transport to the laboratory.  For processing, samples are freeze dried and finely ground, then weighed into tin cups that are pressed to remove air. Samples analysed for d13Corganic are treated with 10% HCl and dried at 50 degrees Celsius before placing into tin cups. Enough sample should be weighed to contain at least 35 ug of nitrogen and 150 ug of carbon. Samples and lab standards are placed into an AS autosampler of an elemental analyser (such as Thermoquest; Carlo Erba Instruments NC 2500).  The sample is dropped from the autosampler into a quartz combustion column set at 1000 degrees Celsius.  This column is packed with chromium oxide, silvered cobaltous oxide, quartz wool and an ash tube.  The introduction of the sample into the combustion column is timed to coincide with a pulse of ultra high purity (UHP) oxygen to combust the sample to form NOx, CO2, and H2O gases and a residual packet of ash.  These gases are transported in a carrier gas of UHP helium to a reduction column where NOx is converted to N2.  The reduction column is set at a temperature of 750 degrees Celsius and is packed with copper wire and quartz wool.

After the reduction column, H2O is removed from the sample gas with a trap of magnesium perchlorate.   Next the N2 and CO2 are separated with a gas chromatograph column.  The remaining gases are carried to an open split of a Finnigan, Conflo III where the sample gas is taken up with a fine glass capillary that leads to the source of a Finnigan, Deltaplus mass spectrometer.  

 (-1.06) for d13C. Nitrogen isotopic values are listed relative to air (vs air) and carbon isotopic values are listed relative to Vienna Peedee Belemnite (vpdb).

More generally, the isotopic composition of organic carbon (d13Corganic) should be determined on decarbonated (10% HCl) subsamples using a VG PRISM isotope ratio mass spectrometer, with a Carlo Erba CHN analyzer fitted in-line as the gas preparation device (Calvert et al., 1995). The isotopic data for organic carbon are reported in the conventional d-notation (equation 2) with respect to the PDB standard.  d15Ntotal values are determined on a second set of untreated subsamples using the same CHN-PRISM setup (Waser, Yin et al. 1998; Waser, Harrison et al. 1998). The results are reported relative to air N2 (equation 3) and the precision should be ±0.3 ppt. An operating system of windows NT 4.0 and ISODAT software (supplied by Finnigan) is used to obtain d15N and d13C isotopic values in the same run.  This value is corrected in an excel spreadsheet using a calibration curve of expected lab standard isotopic values versus measured lab standard isotopic values for various lab standards included in the sample run.  Lab standards are calibrated against the international standards IAEA-N1 (+0.4 per mil) and IAEA-N2 (+20.3 per mil) for d15N and NBS 19 (1.95) and NBS 20. 

The isotopic composition of organic carbon is measured relative to the PDB (Pee Dee Belemnite) standard and is reported as follows:


The isotopic composition of total nitrogen is referenced to air and is reported as follows:


APHA 1998. Standard Methods for the Examination of Water and Wastewater, 20th Edition.

American Public Health Association. Washington, D.C.

Allen HE, Fu G, Boothman W, DiToro D, Mahony JD. 1991. Draft analytical method for determination of acid volatile sulfide in sediment. U.S. Environmental Protection Agency, Washington, DC

Allen HE, Fu GM, Deng BL. 1993. Analysis of acid volatile sulfide and simultaneously extracted metals for the estimation of potential toxicity in aquatic sediments. Environ Toxicol Chem 12:1441–1453.

Bloom, N.S., and Crecelius, E.A., 1983, Determination of mercury in seawater at sub-nanogram per liter levels: Mar. Chemistry, v. 14, p. 49-59.

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Gobeil, C., Macdonald, R.W., Smith, J.N. and Beaudin, L. (2001). Atlantic water flow pathways revealed by lead contamination in Arctic Basin sediments. Science, 293, 1301-1304.

Hammerschmidt, C.R. and Burton, Allen Jr. 2010. Measurements of acid volatile sulfid and simultaneously extracted metals are irreproducible among laboratories. Environmental Toxicology and Chemistry, 29,1453–1456.

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Mortlock, R. A. and P. N. Froelich (1989). “A simple method for the rapid determination of biogenic opal in pelagic marine sediments.” Deep Sea Research 36(9): 1415-1426.

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Verardo, D. J., P. N. Froelich, et al. (1990). “Determination of organic carbon and nitrogen in marine sediments using the Carlo Erba NA-1500 Analyzer.” Deep Sea Research 37(1): 157-165.

Waser, N. A., K. Yin, et al. (1998). Nitrogen isotope fractionation during nitrate, ammonium and urea uptake by marine diatoms and coccolithophores under various conditions of N availability. Marine Ecology Progress Series 169: 29-41.

Waser, N. A. D., P. J. Harrison, et al. (1998). “Nitrogen isotope fractionation during the uptake and assimilation of nitrate, nitrite, ammonium, and urea by a marine diatom.” Limnology and Oceanography 43(2): 215-224.

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