Methodology adopted by Department of Ecology (Procedures and equipment)

Microphytobenthos analysis

Three replicates of 1 cm3 of sediment are collected from sediment cores. The replicates are poured with 10 ml of 2% glutardehyde. To avoid osmotic shock formaldehyde solution was prepared on prefiltered seawater. In laboratory, 0.5 to 1 cm3 of two replicates were diluted in 10 ml of the 2% glutardehyde sollution and settled in sedimenting chambers (10 ml volume). Each of the two replicates is analysed under the inverted microscope. As minimum 500 cells is counted and identified from each replicate. The third replicate was oxidised in 30% of peroxide and mounted on Naphrax for qualitative analyses of diatoms composition.

Phytoplankton & microplankton

Water column

Phytoplankton (microplankton) is collected with the use of perplex glass bottle of 5 dm3 volume with closing device (Niskin Bottle or similar design). Samples are collected from the depths delimited after the light measurement (usually at surface, 50%, 10% and 1% of surface irradiance), in the absence of light meter the samples are collected from 0, 2, 5,10 and 50 m depth. Water sample onboard is placed into 250 cm3 jar, fixed with Lugol solutiona and 3% formaldehyde. Samples are analysed in the lab under the inverted microscope, after the sedimentation in 1 ml chambers. Phytoplankton is counted under the 200 x magnitude, and as minimum 1000 objects are identified and counted from each sample.

Sea ice

Ice samples are collected with the corer (drill) from the surface or by SCUBA diver from below. The ice core of 5 cm diameter is sliced into 5 cm thick parts, each one is placed in separate plastic zip bag. In the laboratory, the ice core is placed in the 250 ml filtered seawater, and left in 4°C in cooler for slow thawing. After the melting is completed, the sample is palced in the jar and fixed with Lugol solution and 3% of formaldehyde. Sorting and counting procedures are the same as for the phytoplankton.

Conical net of 20 mesh size for qualiatative phytoplankton sampling

Cascade filtering device with two different filters mounted; upper filter with 10 µm, lower with 0.4 µm mesh size

Niskin bottles, 5 and 15 l volume



WP2 net with 57 cm diameter, 180 um gauze, closing device and flow meter is the standard net used for vertical hauls.µ Routinely three water strata are sampled, delimited after CTD measurements (usually: surface water - 0-20 m, intermediate layer - 20-50 m, below picnokline - 50-200 m). Vertical sampling in layers thinner than 10 m is not practical. Once retrieved, the net is being washed externally with the hose with seawater onboard, what allows flushing down the plankton attached to the net column. Then the collector is removed and emptied into the small plastic bucket. The collector's walls are rinsed gently to remove all remaining plankton and the sample is poured into the jar of 250 ml volume. Excess water is being filtered out through the same gauze. The sample is preserved with 37% formaldehyde buffered with borax, to obtain 4% formalin solution in the plankton sample. Second standard device is multinet sampler (Hydrobios-Kiel) with 0.25 m2 opening, equipped with 5 nets with 180 µm mesh size and flow meters. Net is hauled vertically through the five discrete water layers, established after the CTD profiling. The sample treatment is the same as described above.


In the laboratory, the jar with zooplankton sample is placed under the ventilation funnel. The sample is gently rinsed with tap water on the 180um gauze sieve to remove the formalin. If needed sample is poured into the splitter and divided into equal parts before analysis. Washed sample is placed into 100-200 ml glass. All the conspicuous and large specimens are removed first and identified. From the remaining plankton, series of five subsamples are taken by 2 ml pipette and put into the glass or plastic Petri dishes with an mm grid on the bottom. Organisms from each sub- sample are identified, counted, and measured under the stereomicroscope. At least 1000 small sized specimens are identified from the sample. The rest of is looked through to check for the large individuals, rare species and macroplankton. Finally, the number of specimens counted is recalculated to the original sample volume. After the identification, plankton is placed back into the original jar.


Multinet - main part of the gear, before mounting the nets

Multinet - colectors are numbered and placed in safe box that prevent falling down

Messenger with safety wire, type used for water casts and plankton nets with closing device

WP2 - plankton is being gently washed from the gauze into sample jar

WP2 - closing device mounted on WP2 net

WP2 mesozooplankton net before lowering

Juday net

WP3 net

Microfauna (Ciliates) in sediments

The sediment core is obtained by inserting 36 x 297 mm plastic core into the sand, closing with a rubber corks. The core is placed in the lab 30 minutes after collection, and sliced in 2 cm sections to 20 cm sediment depth. Subsamples for examination are chosen by inserting into each sediment slice a 12 x 20 mm plastic mini-core, giving a subsample volume of 2,26 ml. The subsamples are fixed in 10 ml 1% glutaraldehyde solution (diluted with filtered water from the sampling l°Cation). The samples are then stored in refregirator in 50 ml plastic containers at 4°C.
    Within two days from subsamples separation, the sediment slices are washed, by shaking and then allowed to stand for approximately 5 s. A 5 ml sample of water is removed with an Eppendorf pipette. The remaining sample is then supplemented with adding 5 ml 1% glutaraldehyde solution, and after shaking another 5 ml of supernatant is removed. The procedure is repeated 5 times, resulting in 25 ml of material from the washed sample. Assuming that ciliates were randomly distributed in the solution immediately after shaking, 5 rinses removed 97% of the ciliates present in the initial sample, because 0,55 x 100 = 3.1% of the original sample remaining.
    After shaking the obtained supernatant 10 ml is placed into plexi glass sedimentation column for at least 6 h. After this time settled ciliates are counted using light microscopy at a magnification of 200 x and 400 x. Then cells are photographed, measured and identificated to the lowest taxonomic level. Abundance is presented in number of specimens per 2.26 ml of sample. The biomass is calculated using the conversion factors from the literature: 0.11 (ed. Edler 1979) and presented in ug C / cm3.
    For the examination of living material, two sediment cores are extracted and sliced as described above, but not fixed. Sediment sections from cores were slipped with minimal disturbance to a 50 ml plastic recipient and were immediately carried to the laboratory. Extraction of the ciliates was made by the sea-water ice method (Uhlig 1965). The cells were counted by removing them one by one with a pipette under the dissection microscope and individual ciliates were photographed using light microscopy at a magnification of 200 x or 400 x. Selected ciliates were stained with protargol according to the method of Wilbert (1975.) (Foissner et al.1999). A mixture of Bouin's fluid with saturated mercuric chloride (1:1) was used as fixative. Photos and durable slides have been used to identify species.

Carey P.G. (1992) Marine Interstitial Ciliates. Chapman & Hall. 1-351.
Edler L.(ed.) (1979) Recommendations on methods for marine biological studies in the Baltic Sea. Phytoplankton and chlorophyll. National Swedish Environment Protection Board. 1-38.
Foissner W., Berger H., Schaumburg J. (1999) Identification and Ecology of Limnetic Plankton Ciliates. Bavarian State Office for Water Management. 1-793.
Uhlig G. (1965) Untersuchungen zur Extraktion der vagilen Mikrofauna aus marinen Sedimenten. Akademische Verlagsgesellschaft Geest & Portig K.-G..151-157.

Meiofauna in sediments

Meiofauna is collected from the sediment cores, obtained from a perplex tubes of 3,6 cm diameter (surface ~10 cm-2 is appropriate for all types of sediment), inserted 15 cm deep into the seabed. From one site six (at least three) replicates are taken.
Sediment is gently pushed out from the tube and cut into the slices. Te slices are usually taken from particular layers: 0-1, 1-2, 2-3, 3-4, 4-5, 5-10 and 10-15 cm. Sometimes core is cut every 1 cm into even slices. Each sediment slice (subsample) is placed into the separate jar, fixed with 4 % neutral formaldehyde solution and stained with Bengal Rose, to obtain a pink/reddish color of the sample.
For extraction of meiofauna from sediment are used two methods depending of the amounts of the detritus or silt-clay in the sediment.
Decantation method - when the sediment is a sand with low amounts of detritus or silt-clay.
In the laboratory, each slice is placed in the 1000 ml cylinder, filled with tap water and shaken vigorously, to suspend the sediment grains. The water is then filtered through 0.038 mm screen, and the procedure of shaking and flotation is repeated 10 times. All meiofauna organisms are retained on the screen, are gently washed into the Petri dish with measuring grid on the bottom and counted under the low power stereo microscope.
Density gradient centrifugation method - the extraction from mud or detritus is most efficiently using a density gradient in a centrifugation procedure (Heip et al. 1985).
In this method liquid with a density larger than the density of meiofaunal organisms can be used (Ludox, density of 1.15).
The method consist: placement of sediment sample into the Ludox solution, centrifuge it with 1800 g for 10 min. The meiofauna organisms are retained in the 'gel', once the sediment is on the end of the tube. Repeat centrifugation three times more. This method does not work for heavy foraminifera, since their density is often equal to the sediment grain.
In the lab, the meiofauna organisms are counted on the measuring grid, usually 1000 specimens per sample (sediment slice).

Basic handboks:
Heip C., Vincx M., Vranken G., 1985, The ecology of marine nematodes. °Ceanographic Marine Biology Annual Review 23: 399-489.
Vincx M., 1996, Meiofauna in marine and freshwater sediments. [In:] Methods for the examination of organisms diversity in soils and sediments (ed G.S. Hall). CAB INTERNATIONAL

Meiofauna sampling - large syringes are inserted into box corer sediment sample


Sesile soft bottom fauna, sublittoral - Van Venn grab, with flap covers, 40 kg weight, opening 30 x 30 cm, triplicate samples from one sampling point. After collecting the sample, first open the grab onboard, using plastic spade remove gently upper 2 cm of sediment into the plastic 5 dm3 bucket, wash gently with filtered seawater and sieve on 0.5 mm screen. Place the remaining sediment into the washing tray, use gently filtered seawater from the hoose and sieve organisms by flotation on 0.5 mm screen. After washing the sample, check large stones and gravel for the presence of sessile organisms, remove clean rocks, and preserve organisms in 4% formaldehyde solution in filtered seawater. In case of substantial amount of sediment left use 10% formaldehyde solution. In the laboratory sample is placed in the plastic container to washout the formaldehyde, and is gently sorted again to remove excess sediment. Organisms remaining on 0.5mm sieve are sorted under the low power stereo microscope. From each sample all macrofauna organisms are sorted, divided into separate jars containing main taxonomic groups. All organisms are counted, for crustaceans the body length is measured for each specimen (tip of rostrum to telson), biomass (wet weight) is established after gently blotting animal (or the whole number of specimens from given taxon) on filter paper and weighted with 0.1 mm accuracy.

Sessile hard bottom fauna, sublittoral - SCUBA diving in the depth of 2 to 30 m, three randomly distributed metal frames (30x30cm) on the selected type of habitat, within the same depth. All organisms are collected (cut from the surface) from each frame into the mesh bag of 0.5 mm size. In case of motile organisms collection form the hard botto m, the frame is equipped with gauze sac. The ejector powered from the air bottle might be used for collection of smaller, unattached animals hidden among rocks and stones.

Motile epibenthos - light epibenthic sledge, equipped with solid fabric apron, thick coarse net and 1mm mesh size gauze in the cod end. The dredge is hauled along the previously recognized bottom profile, within given depth interval, between 2 m and 300 m. Usual ratio of a cable length needed to haul the dredge is 3 times the depth needed, the hauling speed does not exceed 2 knots, and a haul duration range from 10 to 30 minutes. The whole dredge is hauled out on the working deck, and depending on the type of sample obtained (less or more muddy) a subsample or the whole macrofauna is hand picked. The sample is a qualitative one, giving only the rough proportion of the density of organisms living on the seabed. Having number of dredge samples along a transect, permits to get the good information on the macro faunal species richness from the analysed area.

Carrion feeders - number of fast moving organisms may avoid grabs and dredges or are too small to be collected into the fishing gear like beam or otter trawl. Carrion feeding crustaceans belong to the group of animals that are best collected with the use of baited traps. The one used in our department is a cylindrical construction of 1m per 30 cm, covered with 1mm gauze, with two conical entrances on both ends. The bait (100 g piece of fish or bird’s meat) is placed in nylon mesh bag inside the trap. Trap is lowered on the seabed with an 2 kg weight, and fastened to the buoy on the surface. Operational depths are 2 to 200 m, deeper on, the acoustic release system is practical. The trap is exposed from 2 hours up to 24 hours, and retrieved onboard. The catch (may reach over 1 kg of biomass) is removed from the opening on the side of the net, washed in filtered seawater and placed in the preservative. The position, amount of bait before and after the exposure, depth, duration of exposure are noted.

Intertidal sedimentary bottom - quantitative sampling is performed with the use of plastic tubes of 10 cm diameter, inserted at least 10 cm into the sediment. Three to six replicates are collected from any site, usually organized along the water marks - at Low Water, Mean Water and High Water Mark. The coarser the sediment the deeper the sample should be collected (in mud the upper 10 cm, in coarse sand the upper 40 cm).

Intertidal hard bottom - quantitative sampling is performed with the use of 20x20 cm metal frame, placed on the selected water mark during ebb tide. All organisms are cut, scraped from the frame area and gently collected into the collector's jars. Three to six replicates are recommended.

Basic handboks:
Holme AN, Mc Intyre AD. 1971. Methods for the study of marine benthos. IBP Handbook no 16, Blackewll Sci. Publ. Oxford, 334 pp

Van Venn grab

Container for washing sediment samples

Small box corer

Epibenthic sledge

Camera and video recording system ready for lowering

Epibenthic sledge

Box corer - removable box with newly collected sediment

Sedimentation measurements

Suspensions measurements

Water samples of 1.7 dm-3 are collected with the water sampler (Niskin bottle type) from various depths, depending on the sampling design (usually 0 m, 5 m, 50 m depth). Water subsamples of 100 to 500 ml are filtered on combusted and pre weighted Whatman Glass Microfibre filters GF / F, 0.7 µ to get concentration of Particulate Total Mater (PTM). Further procedures as described above with sedimentation samples processing.
Gravity flows are measured one meter over the sea bottom with Sensordata currentmeter Mini SD 6000 combined with Seapoint Turbidity Meter emitting light 880 nm and scatterance angles 15-150 degrees. Empirical data of solids concentration from Adventfjorden are then used to calibrate backscatter by plotting PTM (mg dm-3) against FTU (Formazin Turbidity Units) recorded by Seapoit Turbidity Meter. Received linear regression equation TPM = 0.9986 FTU + 0.6899 (determination coefficient R2 = 0.999) was used to calculate concentration of PTM and gravity flows.

Sedimentation measurements

The sediment traps are cylinders of perplex, 9.4 cm in diameter and 70 cm in length or 10 / 100 cm.. Equipped with the outlet 100 cm from the bottom. Traps are deployed vertically in the water column, attached to the rope with buoy in such way, that allow them to hang vertically (Zajączkowski 2002) Traps are deposited from 12 to 48 hours, depending on the sedimentation intensity. Traps are deployed below euphotic zone or below the pyknokline, 1m above the seabed and usually in between in water column. Once retrieved, the samples is placed onboard vertically, to avoid mixing the collected sediment, and the excess water is removed through the outlet. Remaining 1 dm-3 of water with the accumulated sediment is placed into the sampling jar. In case of delay between collection and lab processing the sediment sample is fixed with 3% of formaldehyde.

In the lab, suspensions subsamples are filtered in combusted and pre-weighted Whatman Glass Microfibre filters GF/F 0,7 µm, usually 100 to 200 ml are filtered. After the filtration, the distilled water is used to wash out the salt particles from the filter. Some filters are kept frozen at -18° C for further analyses (maximum 12 days storage for chlorophyll a), filters meant for the TSM (Total Sedimented Matter ) analyze are dried in 60° C to stable weight, weighted, and next burned in 450° C for 24 hours to combust the organic particles. After the burning, the filters are weighted again, and the remaining weight is considered as ISM (Inorganic Sedimented Matter). The organic matter amount is calculated as a weight loss in combustion. Additionally, subsamples of 50 to 100 ml of water are filtered on combusted filters for particulate organic carbon (POC) and nitrogen (PON) analyses. Before processing on Perkin Elmer 2400 CHN analyzer, samples are stored 24 h in vapours of hydrochloric acid to remove of calcium carbonate, and then ventilated 8 h (Hadges, Stern 1984). Part of the original suspensions sample (usually 50 ml) is placed in the glass cylinder for 24 hours for settlement of suspensions, and after removing the excess water sediment sample is examined under the microscope for particles indentification.

Hadges, J.I., Stern J.H. 1984, Carbon and nitrogen determinations of carbonate containing solids. Limnol. oceanogr. Vol. 29: 657-663.
Zajaczkowski M., 2002, On the use of sediment traps in sedimentation measurements in glaciated fjords. Polish Polar Research. Vol. 23: 161-174.

The double sediment trap

The current meter net

Kullenberg piston corer, 3 m long section onboard

Kullenberg piston corer ready for lowering

Inner part of the Kullenberg piston corer, plastic PVC pipe containing sediment is being cut to sections

Niemisto sediment corer ready for lowering

Suspensions measurements

Water samples of 1 dm3 are collected with the water sampler (Niskin bottle type) from various depths, depending on the sampling design (usually 0 m, 5 m, 50 m depth). Water subsamples of 100 to 500 ml are filtered on pre weighted GFC filters of 045 µm. Further procedures as described above with sedimentation samples processing.

Sediment analyses

  1. Granulometry table with Wenworth scale of sediment size.

    The sieves set with shaker
  2. Organic matter content.
    Organic matter content is measure in sediment samples of 1g dry mass, previously dried in 65°C to stabilize weight. Sample is placed in the marked, pre-weighted china jar in oven and burnt in 450° C for 24 hours. After the cooling of the oven, the sample is weighted again in the jar, and the weight loss is considered to be the organic matter loss in combustion. The organic matter content is presented as the % of original dry sediment weight. Organic Carbon Content in sediment - the samples of 1 g dry sediment are analysed.
  3. Porosity.
    Porosity represents the amount of water that may fill the given sediment unit (mm3 of water per g of sediment)
  4. Permeability.
    Permeability presents the rate of water penetration through the sediment and is expressed in mm3 per unit of length (cm) in time (sec).

    The equipment set for the permeability measurements
  5. Chlorophyll a content Sand sample (surface 0.5 cm of sand thickness, and 2 cm in diameter) is placed in the glass jar filled with 25 ml of distilled water, shaken vigorously for 15 minutes, and filtered through the GFC Whatman cellulose filter. Filter is folded in half and frozen in -18°C, for no more than 14 days, within that time the another extraction in acetone is performed, and the supernatant is analysed in spectrophotometer 460 to 670 nm wavelength. Three sub-samples are taken for spectroscopic meaurments.

Pore water sampling

Pore water shall be removed from distinct sediment layer, avoiding contamination of sample from the water column. For the sediment depths up to 30 cm, different kinds of syringes are used, equipped with screen that protect surface water from penetrating the sediment. For deeper sediment (30-60 cm) the peeper is used - double steel tube with number of small holes along its length. Surface, fresh water outflows is collected with the Bokuniewicz method - a barrel or tray with attached plastic bag is digged into the sand and after 1-6 hours it collects the emiting freshwater.

Carbohydrate concentration and composition (for sandy sediments)

I. Sampling:
  1. Collect one sediment core (16 cm length, 3.6 cm diameter) from each following depths: shore line, 1.5 m and 7 m
  2. Slice each sediment core for 8 slices (samples): 0-2 cm, 2-4 cm, 4-6 cm, 6-8 cm, 8-10 cm, 10-12 cm, 12-14 cm, 14-16 cm
  3. Freeze samples
II. Total Carbohydrate measurements:
  1. Weight out 0,5g of each samples into test tubes. Record exact weight of each sub-sample. For each sample should be weight out 3 sub-samples
  2. Add to each sub-samples 1 ml of dH2O
  3. Prepare 3 blank: add to empty test tubes 1 ml of dH2O
  4. Add to each test tube 0.5 ml of 5% phenol then quickly add 2.5 ml of conc. H2SO4, ensuring that the sub-samples are well mixed
  5. Leave to cool for ~ 30 min
  6. Read in spectrophotometer at 485 nm
III. Colloidal carbohydrate measurements:
  1. Weight out 1.0 g of samples into centrifuge tubes. Record exact weight of each sub-sample. For each sample should be weight out 3 sub-samples
  2. Prepare 3 blank: add to empty centrifuge tubes 1 ml of dH2O
  3. Add to each centrifuge tube 4 ml of 100 Mm EDTA and shake in a vortex mixer
  4. Leave in a water bath at 25 C for 30 min
  5. Centrifuge for 15 min at 4000 g
  6. Transfer 1 ml of supernatant to test tubes (each sub-samples)
  7. Add to each test tube 0.5 ml of 5% phenol then quickly add 2.5 ml of conc. H2SO4, ensuring that the sub-samples are well mixed
  8. Leave to cool for ~ 30 min
  9. Read in spectrophotometer at 485 nm
IV. EPS carbohydrate measurements:
  1. Transfer 1.5 ml of supernatant from the colloidal fraction to centrifuge tubes (each sub-samples and blanks)
  2. Make up each of them to 70% ethanol (add 3.5 ml industrial methylated spirit)
  3. Leave overnight to precipitate at 4 °C (cover tubes to minimize loss)
  4. Centrifuge at 4000 g for 15 min
  5. Remove all supernatant and re-suspend EPS pellet in 1 ml of dH2O
  6. Transfer 1 ml of each re-suspended EPS to test tubes (blanks too)
  7. Add to each test tube 0.5 ml of 5% phenol then quickly add 2.5 ml of conc. H2SO4, ensuring that the sub-samples are well mixed
  8. Leave to cool for ~ 30 min
  9. Read in spectrophotometer at 485 nm
V. Bounded carbohydrate measurement [with dH2O]:
  1. Add to each of the pellet from the colloidal fraction 4 ml of conc. ethanol (blank too)
  2. Shake in a vortex mixer
  3. Centrifuge at 4000 g for 15 min
  4. Remove all supernatant (if it is colored: even few times - until supernatant will be colorless)
  5. Add 4 ml of dH2O and shake in a vortex mixer
  6. Leave in water bath at 95 C for 1 h, shake in a vortex mixer after 30 min of bath
  7. Centrifuge at 4000 g for 15 min
  8. Transfer 1 ml of supernatant to test tubes (remove rest of supernatant)
  9. Add to each test tube 0.5 ml of 5% phenol then quickly add 2.5 ml of conc. H2SO4, ensuring that the sub-samples are well mixed
  10. Leave to cool for ~ 30 min
  11. Read in spectrophotometer at 485 nm
VI. Bounded carbohydrate measurement [with NaHCO3]:
  1. Add to each pellet (bounded with dH2O) 4 ml of 0.5 M NaHCO3 and shake in a vortex mixer
  2. Leave in water bath at 95 C for 1h, shake in a vortex mixer after 30 min of bath
  3. Centrifuge at 4000 g for 15 min
  4. Transfer 1 ml of supernatant to test tubes (remove rest of supernatant)
  5. Add to each test tube 0.5 ml of 5% phenol then quickly add 2.5 ml of conc. H2SO4, ensuring that the sub-samples are well mixed
  6. Leave to cool for ~ 30 min
  7. Read in spectrophotometer at 485 nm
VII. Standard Curve:
  1. Dissolve 1g anhydrous glucose in 100 ml of dH2O (10 mg / ml)
  2. Take 1 ml of 10 mg / ml glucose solution and make up to 100 ml with dH2O (100 µg / ml)
  3. Make 7 standard solutions:
    a) 0 ml of 100 µg / ml glucose solution + 10 ml of dH2O (0 µg / ml)
    b) 1 ml of 100 µg / ml glucose solution + 9 ml of dH2O (10 µg / ml)
    c) 2 ml of 100 µg / ml glucose solution + 8 ml of dH2O (20 µg / ml)
    d) 4 ml of 100 µg / ml glucose solution + 6 ml of dH2O (40 µg / ml)
    e) 6 ml of 100 µg / ml glucose solution + 4 ml of dH2O (60 µg / ml)
    f) 8 ml of 100 µg / ml glucose solution + 2 ml of dH2O (80 µg / ml)
    g) 10 ml of 100 µg / ml glucose solution + 0 ml of dH2O (100 µg / ml)
  4. Transfer 1 ml of each solution to test tubes (3 samples for each solution)
  5. Prepare 3 blank: add to empty test tubes 1 ml of dH2O
  6. Add to each test tube 0.5 ml of 5% phenol then quickly add 2.5 ml of conc. H2SO4, ensuring that the samples are well mixed
  7. Leave to cool for ~ 30 min
  8. Read in spectrophotometer at 485 nm

Safety onboard
Always be careful and wear safety clothes. Never ever drink alcohol onboard research vessel (at least when working).
In Arctic be aware of wild animals - especially polar bears.
Safety First! Brightly colored jacket and helmet are a must while operating heavy gear onboard.