Functional Groups Collected
Sediments
Pool top 2cm from 5, 60 ml syringe cores, put in 4 oz Qorpack jar and dry @ 50o C
Benthic diatoms
Nitex migration, at low tide lay down 210u Nitex on sediment surface, wait 20 min for diatoms to migrate, rinse diatoms onto GFF at lab, dry @ 50o C
Nereis
Using shovels, dig up15-20, 2-5 cm lengths, place in jar with ambient water to purge guts for 24 hr, dry @ 50o C
Mummichog
Using seine, collect 15-20, 35-50 mm lengths, fillet and save flesh tissue (tail end), dry @ 50o C
Ribbed mussel
Pool 15-20, below S. alt. on main stem, abductor muscle, dry @ 50o C
POM
mid ebb, water column water filtered, clogged on GFF, dry @ 50o C
Blue mussel
Pool 15-20, on front edge of main stem, abductor muscle, dry @ 50o C
Pelagic copepod
mid ebb, Plankton net, 150 u, light migration, GFF, dry @ 50o C
Silverside
Seine, pool 15-20, flesh tissue (tail end) like mummichogs, dry @ 50o C
Mya (softshell clam)
Pool 15-20 from tidal flats, abductor muscle, dry @ 50o C
Bivalves (all species)
Pool 15-20, measure length, abductor muscle, rinse in DI H2O, dry @ 50o C
Marsh plants (species will vary in estuary)
Aboveground biomass, green tissue, cut into 5” pieces, soak in bucket of tap water 5 minutes, rinse with DI H2O, dry @ 50o C
Other fish species
From seine, flesh tissue, rinse in DI H2O, dry @ 50o C
Stable Isotope Food Web Monitoring Sample Collection and Processing
Sediments
Collect 5 surface (2cm) cores using 60 ml syringe, place cores in a labeled (date, station, sample type) 4 oz Qorpack jar and in a cooler for transport back to the lab. Freeze if the sample cannot be dried soon.
Dry sediments @ 50o C. After the sample is dried, use a dissecting microscope and check sediment for carbonates using a few drops of 10% HCl on a small subsample, if bubbling occurs then a larger subsample should be acidified and redried for isotope analysis. If carbonates exist it is necessary to remove them, as they will interfere with the 13C analysis. Grind a subsample of the sediments (Wig-L-Bug) for stable isotope analysis and place in a clean scint vial.
Benthic diatoms
Collect benthic diatoms by laying down several (5-7) separate pieces of 210u Nitex on the sediment surface at low tide. Pick a sediment surface that has a golden sheen and still looks fairly wet. If the surface is wet enough the diatoms will migrate up through the Nitex and there will be a fairly pure sample on one side of the Nitex. Use a 500 ml squirt bottle filled with ambient water from the site and gently squirt drops onto the Nitex to get it to lay on the sediment surface without air gaps. After 20 minutes or so, peel the Nitex from the sediment surface and check for an adequate golden sheen on the Nitex, be careful not to contaminate the diatom side with the under side mud or sand. Fold the Nitex (diatom side) carefully in on itself and place in a labeled (date, station, sample type) ziplock bag and put bag in a cooler for transport back to the lab.
At the lab unfold the Nitex with the diatom side on the outside, fold it so that the middle of the Nitex forms a corner facing down and carefully using the ambient water squirt bottle, rinse the diatoms off onto an ashed 25 mm GFF filter that has been set up in a 25mm Gelman filter tower and flask (need replicate filters). When the filter/liquid looks green suction it dry and place the filters into separate labeled (date, station, sample type) small petri dishes. Put petri dishes in a freezer if the samples cannot be dried soon. Dry the filters @ 50o C. After the sample is dried, use a dissecting microscope and check the filters for carbonates using a drop of 10% HCl on a portion of the filter, if bubbling occurs then the whole filter should be acidified and re-dried for isotope analysis. If carbonates exist it is necessary to remove them, as they will interfere with the 13C analysis.
Nereis
Collect Nereis worms by digging with a shovel in the tidal flat sediments; often the peat chunks have more worms. Collect 15 to 20 worms of 2-5 cm lengths and place in a jar with ambient water. Place the jar in a cooler for transport back to the lab.
At the lab transfer the worms to another jar and put clean ambient water in the jar, let the worms sit overnight to purge their guts, then count and record the lengths of worms, towel blot and place worms in a labeled (date, station, sample type) glass scint vial and freeze. Dry the worms @ 50o C and then grind the worms (Wig-L-Bug) for stable isotope analysis and place back in the labeled (date, station, sample type) scint vial.
Mummichog
Using a seine collect 15 to 20 mummichogs, 35-50 mm lengths and place in a labeled (date, station, sample type) ziplock bag and place in cooler for transport back to the lab. At the lab, count and record the lengths of the mummichogs, fillet and save tissue in a labeled (date, station, sample type) glass scint vial and freeze. Dry the mummichogs @ 50o C and then grind the mummichogs (Wig-L-Bug) for stable isotope analysis and place back in the labeled (date, station, sample type) scint vial.
Ribbed mussel
Collect 15 to 20 ribbed mussels from below the Spartina alterniflora channel bank edge and place in a labeled (date, station, sample type) ziplock bag and place in cooler for transport back to the lab.
At the lab, count and record the lengths of the mussels, cut out the abductor muscle and rinse/dip in DI water. Save muscle tissue in a labeled (date, station, sample type) glass scint vial and freeze. Be careful not to get shell fragments mixed in with the abductor muscle. Dry the tissue @ 50o C and then grind the tissue (Wig-L-Bug) for stable isotope analysis and place back in the labeled (date, station, sample type) scint vial.
Particulate Organic Matter (POM)
Collect between 1 liter and 4 liters (depends on station) of mid ebb channel water into labeled (date, station, sample type) bottles recording date and time of day, place in cooler for transport back to the lab.
At the lab, shake the bottles thoroughly and filter water onto ashed, 25 mm GFF filters until clogged, rinse with DI H2O (need replicates) and place filters into separate labeled (date, station, sample type) small petri dishes. Put petri dishes in a freezer if the samples cannot be dried soon. Dry the tissue @ 50o C.
Plankton Tow 20u (POM, 20u for 34S)
Use 20u plankton net and either pour buckets or tow net (tow slowly or net will tear) to get noticeable amount of material for 34S analysis. POM GFF filters do not have enough mass for 34S analysis. Transfer out of cod end into labeled jar. In lab, concentrate on 20u Nitex, 3 rinses with DI H2O (3x the volume of sample) and transfer to labeled scint vial and freeze. Dry @ 50o C, check particle size before submitting for analysis.
Blue mussel
Collect 15 to 20 blue mussels from the front edge of the channel and place in a labeled (date, station, sample type) ziplock bag and place in cooler for transport back to the lab.
At the lab, count and record the lengths of the mussels, cut out the abductor muscle and rinse/dip in DI water. Save muscle tissue in a labeled (date, station, sample type) glass scint vial and freeze. Be careful not to get shell fragments mixed in with the abductor muscle. Dry the tissue @ 50o C and then grind the tissue (Wig-L-Bug) for stable isotope analysis and place back in the labeled (date, station, sample type) scint vial.
Zooplankton (pelagic copepods)
Collect zooplankton from mid channel at mid ebb tide using 150u plankton net. Pour cod end sample into a labeled (date, station, sample type) wide mouth jar, recording date and time of day, place in cooler for transport back the lab. Don’t let jar overheat, the zooplankton must be kept alive.
At the lab, we will conduct a zooplankton light migration technique procedure to purify the sample. Place the jar on a counter and let it settle. When you see a lot of zooplankton in the water column, pour off the water and zooplankton into two glass 250 ml graduated cylinders that have been wrapped with tin foil within 3” of the top. The water height should be about 2 cm above the height of tin foil. Aim a dissecting scope light or flash light at the water surface and let the zooplankton migrate to the light. Set up a vacuum flask for 25 mm filter rig. When there is a dense mass of clean zooplankton at the surface, while avoiding detritus, pipette the zooplankton onto ashed, 25 mm GFF filters (need replicates), squirt filter surface occasionally to migrate zooplankton to the middle of the filter and place filters into separate labeled (date, station, sample type) small petri dishes. Put the petri dishes in a freezer if the samples cannot be dried soon. Dry the filters @ 50o C.
Silversides
Using a seine collect 45 to 60 silversides, 30 – 70 mm lengths and place in a labeled (date, station, sample type) ziplock bag and place in cooler for transport back to the lab. At the lab separate the silversides into 3 subsamples, then count and record the lengths of the silversides for each subsample, then fillet and save the tissue in three separately labeled (date, station, sample type) glass scint vials and freeze. Dry the silversides @ 50o C and then grind them (Wig-L-Bug) for stable isotope analysis and place back in the labeled (date, station, sample type) scint vial.
Mya (softshell clam)
Collect 15 to 20 soft shell clams by digging in the tidal flats and place in a labeled (date, station, sample type) ziplock bag and place in cooler for transport back to the lab. At the lab, count and record the lengths of the clams, cut out the abductor muscle, and rinse/dip in DI water. Save muscle tissue in a labeled (date, station, sample type) glass scint vial and freeze. Be careful not to get shell fragments mixed in with the abductor muscle. Dry the tissue @ 50o C and then grind the tissue (Wig-L-Bug) for stable isotope analysis and place back in the labeled (date, station, sample type) scint vial.
Marsh plants
Above ground biomass, want fresh green leaves. Use anvil clippers to clip stalks/stems, place biomass in plastic bag. Sort plants, separating out green leaves and different species that may be mixed in (S. patens vs Distichlis). Rinse/soak leaves in bucket of tap water to remove sea salt SO4, then rinse with DI H2O. Air dry leaves on bench then place into labeled paper bag with Date, Station, Species and analyses needed. Place paper bag in large drying oven @ 50o C for a few days. Grind plant tissue for stable isotope analysis (Wiley Mill or Wig-L bug check with Marshall Otter for particle size).
Other Organisms
Collect other organisms/species as deemed necessary and follow the above methodology/logic of saving meat/tissue without shell and rinsing adequately with DI H2O to remove seawater sulfate. Care should also be taken to NOT include guts with the tissue samples.
Field Gear and Field lab supplies
Jon boat, 25 HP Honda
150 u plankton net with cod end
Cooler with ice
Dry cooler
Field notebook
Pencils
Sharpies (fine and large)
Label tape
3 buckets
thermometer
Box of 1 gallon ziplock bags
Box of 4 oz Qorpack jars, minimum of 6 jars
2, 60 ml syringe cores
5, 6” x 6” squares of 210u Nitex
500 ml squirt bottle with fine nozzle
shovel
small seine
4, one liter bottles for POM
4, one liter wide mouth Nalgene bottles for zooplankton tows
box of ashed 25 mm GFF filters
filter forceps
small petri dishes for 25 mm filters
hand vacuum pump/flask and Gelman tower set up for 25 mm filters
dissecting kit with scalpels and tweezers
flat of glass scint vials with caps
tin foil
2, 250 ml glass graduated cylinders
2, dissecting scope lights or flash lights (with batteries)
