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6-SAMPLING METHODS FOR ZOOPLANKTON

Zooplankton is typically collected with a fine mesh net, but using buckets or dip nets around bright lights is also possible. The plankton nets used are of various sizes and types. The different nets can broadly be put into two categories, the open type used mainly for horizontal and oblique hauls and the closed nets with messengers for collecting vertical samples from desired depths.

Despite minor variations, the plankton net is conical in shape and consists of ring (rigid/flexible and round/square), the filtering cone and the collecting bucket for collection of organisms (Fig. 2). The collecting bucket should be strong and easy to remove from the net. The netting of the filtering cone is made of silk, nylon or other synthetic material. The material should be durable with accurate and fixed pore size. The mesh should be square and aperture uniform. The mesh size of the netting material will influence the type of zooplankton collected by a net. The nets with finer mesh will capture smaller organisms, larval stages and eggs of planktonic forms and fish eggs while those with coarse netting material are used for collecting bigger plankton and fish larvae. There is a great variety of mesh available from the finest to the coarse pore sizes. In addition to the mesh size, the type, length and mouth area of the net, towing speed, time of collection and type of haul will determine the quality and quantity of zooplankton collected.

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Mesh size, extrusion and avoidance The appropriateness of mesh size can be determined through the trade-off between the net avoidance of zooplankton and net extrusion of zooplankton. With towed plankton nets, the smallest mesh size will never sample all the zooplankton, because larger and better swimming zooplankton will sense the pressure wave in front of a small mesh net and dodge it (this is known as net avoidance). If you use larger mesh, then the smaller zooplankton will be extruded through the mesh. Collection time and speed The zooplankton collections can be made by horizontal, oblique and vertical hauls. In the horizontal sampling the net is towed at a slow speed usually for 5 to 10 minutes. The towing speed of the net should be such that the maximum amount of water enters through the mouth of the net for better filteration and gear used can withstand the strain. The towing speed of the net recommended for horizontal samples is 1.5 to 2.0 knots– or 1 to 2 metres per second–. Any faster will increase the extent of extrusion, and any slower may increase the incidence of avoidance. The net may also be damaged at high speed towing. When the currents are strong, depressors are used to keep the nets in desired position. Type of haul • The horizontal collections are mostly carried out for the surface and subsurface

layers. • In oblique hauls, the net is usually towed above the bottom. • The vertical haul is made to sample the water column. The net is lowered to the

desired depth and hauled slowly upwards. • Closing mechanisms are used to close nets to study zooplankton abundance at

different depths. The choice of net and type of haul to be taken should be determined by the objectives of the study. Examples • The microzooplankton samples can be collected using a 20 µm mesh net, with 20

cm mouth diameter, by vertical or oblique tows. The plankton net is towed for 1-5 minutes depending on the density of plankton.

• The mesozooplankton samples can be collected by oblique tows using a 110 µm mesh net, with mouth opening of 50 cm diameter. The duration of the tows ranged from 1-5 minutes depending on the abundance of plankton at the sampling location.

Whatever type of net is used for sampling, it should be thoroughly washed after each tow so that any planktonic material adhering to the mesh of the filtering cone or other part of the plankton net should be pushed into the collecting bucket to prevent contamination of samples with collections from the previous hauls. The washing of the nets will also prevent clogging especially when there is a bloom or the finer mesh is used for obtaining the samples. The nets should also be checked for torns or holes through which the plankton can escape resulting in the loss of sample.

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After each haul the zooplankton sample is transferred into a cleaned and dried glass beaker of half to one liter capacity. The debris or extraneous material should be removed. Replicate hauls are made whenever possible. For quantitative plankton sampling (to then determine the number or biomass of zooplankton per cubic meter) it is imperative to know the actual volume of water passed through the net. For this purpose, an instrument called flow meter (Fig. 3) is used. It should not be mistaken for current meter. The flow meter has a multi bladed propeller, which is rotated by the flow of water. There is a counter, which records the number of revolutions. The flow meter should be positioned in such a way so that it records the actual flow of water passing through the net. The volume of the water filtered is normally expressed in cubic meters.

Most of the zooplankters migrate vertically in response to light conditions. Their occurrence is poor in upper layers during daytime. For better quantitative and qualitative zooplankton collections, the suitable time for horizontal zooplankton sampling would be before dawn, after dusk or night. Records of the sampling procedures, prevailing environmental conditions and other information should be maintained in the field log sheet. Observation on live plankton could be made in the field for their colouration, abundance and composition prior to fixation and subsequently analysis in the laboratory. The samples should be transported with care otherwise their durability and usefulness would be seriously jeopardized. Fixation After the sampling, the fixation of samples should be carried out, as early as possible, at least within 5 minutes after the collection to avoid damage to animal tissue by bacterial action and autolysis. A fixative actually "fixes" a specimen by stabilizing the proteins within its tissues such that long afterwards, the tissues will still retain a semblance of their appearance in life. An ideal fixative should be cheap and which kills the animals quickly. Again it should be non-corrosive or toxic in nature. The most common fixing and preserving reagent is (4-5%) buffered formaldehyde. It is the cheapest fixative and the zooplankton samples can be stored for number of years. The volume of plankton to solution should be about 1:9. The resulting solution is acidic and therefore is detrimental for specimens containing CaCO3, which unfortunately is present in the skeletons of many taxa. To neutralize or "buffer" the

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acid content, sodium hydroxide, sodium tetraborate, sodium carbonate (NaCO3) or hexamethyene teteramine are sometimes added. The buffers are added in an amount of 200 g to one litre of concentrated formalin. However, this buffering is apparently temporary, as buffered formalin will become acidic again some time after buffering; specimens should never be stored in formalin for extended periods of time. The fixative usually renders the zooplankton body tissues hard and brittle. The additives viz. propylene phenoxetal and propylene glycerol (2 to 5 %) are added to fixatives for flexibility of specimens, resistance to bacteria and moulds. Preservation In contrast to a fixative, a preservative is a solution in which the specimen can be stored and maintained without further degradation. The primary role of the preservative is to provide an environment in which bacterial and other infestations and contaminations are unlikely, thus maintaining the specimens over long periods of time. • Allow 10 days as the minimum fixation periods. • Following fixation, the specimen should be thoroughly but gently rinsed off the

formaldehyde solution in fresh water or distilled water, after which it is usually transferred to preservative for long-term storage.

• The zooplankton are stored in airtight containers with sufficient quantity of preservative.

• Various types of preservatives are available. • The buffered formalin (4 to 5%) is mostly used both as fixative and as the

preservative. The other preservative used is 70%-75% ethanol or 40% isopropanol.

• Glycerin is often added to formalin to prevent shrinkage of specimens, drying of the material and to facilitate retaining colours of zooplankters.

• For better shelf life of the zooplankton samples, the preservative should be changed within the first 6 months.

• It would be better to store the preserved zooplankton samples in well ventilated room at temperature less than 25°C.

• The samples should be kept in the wide mouth glass jars. • A good quality preprinted labels, on which the collector’s name, fixative and

preservative used and other field information are written should be put into the jars for ready reference at the time of sample analysis.

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Zooplankton Diversity and Taxonomy

The zooplankton are very diverse. You should observe your live and preserved samples for zooplankton. The zooplankton samples are analyzed for estimation of biomass (standing stock), enumeration of common taxa and their species. Data obtained is utilized for computation of productivity and faunal and species biodiversity of the study area or ecosystem. Materials: • Dissecting microscopes (1 per student) • Live zooplankton sample • Preserved zooplankton sample • Zooplankton Guide (you can make your own using zooplankton groups or species

that are common to the area where your samples are from) • Computers with internet access (if using online zooplankton guide) • Sea water (if using live marine zooplankton) • Petri dishes

Procedure:

1. Set up your dissecting microscope. Make sure that your light source works and that you know how to focus the microscope. Ask your instructor if you are having trouble.

A. Live Zooplankton sample observation 2. Obtain a live zooplankton sample from your instructor. 3. Use an eye dropper to collect a few drops of the sample and place in a petri dish. 4. Observe the sample with a dissecting microscope. Since the plankton can move

up and down in the drop, you will need to refocus your microscope to see plankton at different levels.

5. Remember to turn off the light when you are not looking in the microscope so that the organisms do not become overheated.

6. Use this sample to study the movements and swimming behavior of different zooplankton forms. Observe the locomotion by antennae in copepods and by cilia in trochophora and veliger larvae.

7. You may want to prepare a sample using methyl cellulose instead of water. This will slow them down (because the methyl cellulose is sticky) and make them easier to see.

8. Describe the live zooplankton sample. List at least 3 things that you notice about the sample.

9. Draw and identify some types of organisms that you viewed in the live sample. Use the zooplankton guide given below to help you identify the organisms.

10. Try to put effort and patience into your drawings and document as many details as you can observe. Take your time to observe structural details, and try to include internal structures (e.g. gut tract).

11. Observe your sample for the following: a. most abundant organisms b. variations in shape, color, and swimming ability c. types of appendages seen on various plankton d. eggs e. larval and juvenile forms of crustaceans and fish

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Swimming responses of zooplankton a. Phototaxis Many plankton are positively phototaxic (drawn to the light). Fill a cylinder with sea water and then introduce about 25 ml of a well-mixed plankton sample. Place the cylinder into a cardboard box with a flashlight at the top. After 2 minutes open the side panel and observe the position of the plankton. Now, shine the light into the bottom of the cylinder. After two minutes, observe the position of the plankton. How can you explain the results Now, let’s investigate the effect of wavelength on this phototaxis. Using red, green and blue colored plastic, test the phototaxis of your sample. Record your results in Table 1. In this table a positive response (+) means the animals were drawn to the light.

Table 1. Response of zooplankton to light

What conclusions can you draw from the results of this experiment? b. Swimming speed Is it possible that these animals could swim enough to account for the vertical migrations seen throughout the day? What is the actual swimming speed of these animals? Using the same cylinder with the light coming from the top and a stop watch, record the swimming speed of ten of the most active animals (most likely animal is copepod). You will need to place a ruler along the cylinder to measure the distance traveled. Compute the average speed. Record the data in Table 2.

Table 2. Swimming speeds of zooplankton

Now, using this speed, compute the maximum distance that could be covered in a 5 hour period (be sure to convert this into meters). B. Preserved Zooplankton Preserved zooplankton samples fixed with 1% (final conc.) formaldehyde will be provided by the instructor for this exercise. Avoid inhaling the formaldehyde (harmful!) and keep sample bottles closed if not in use. You will use the provided plankton samples to study the diversity of zooplankton and to prepare detailed drawings of representatives of the different groups of zooplankton. Details are given below. Include your sheets of drawings in your lab report. Your lab exercise should comprise at least 6 different species. But take the chance to observe many more different species form the samples.

1. Obtain a petri dish of preserved zooplankton from your instructor. Examine the sample under the microscope.

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2. Describe the preserved zooplankton sample. Compare and contrast it to the live zooplankton sample.

3. Draw and identify at least 4 different types of organisms that you viewed in the preserved sample. Use the zooplankton guide to help you identify the organisms.

4. Summarize your findings in a written report.

5. Many of the organisms are too small to be seen with the dissecting microscope. You can prepare a slide of the sample and observe it under a compound scope with low and high power objectives.

For a detailed study and preparation of drawings and documentation, use the plastic Pasteur pipette to capture your specimens of interest while observing through the stereomicroscope. You will see the tip of your pipette; once you have focused the microscope on your specimen of interest, use your second hand to hold the other arm, thereby providing stability to your pipetting hand. After you have captured your specimen from the petri dish, place it with a little drop of seawater (not too much) on a microscope slide, cover with a cover slip (never press the cover slip down, it will smash your specimen), place slide on the compound microscope and commence your observation and drawing. Prepare a detailed drawing of one representative, well-preserved specimen. Next to the drawing, include the zooplankton systematic group name and some features that characterize this group and distinguish it form others. Remember that any drawing without correct and detailed annotations is worth nothing. For the larval forms, note which adult animal belongs to your specimen! Zooplankton can be identified and counted as far as possible with available keys and descriptions until species level. Some specific keys were also be used (Table 3).

6. Repeat this procedure with as many different organisms as time permits.

Table 3: Specific zooplankton identification keys.

C. Prepared sample observations Using your microscope, observe and sketch examples of the following: • Radiolaria These single-celled animals have skeletal elements made of silica. • Foraminifera or Globigerina. Globigerina is one of the most common “forams.”

These animals have shells made from calcium carbonate.

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Pictorial key of zooplankton (Not scale, number in brackets are part number of the guide)

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Detailed Pictorial key of zooplankton (with scale bar)

Fig. 1 Protozoa Ciliata: (1) Vorticella marina, (2a) Zoothamnion marinum, (2b) at a reduced scale showing colonies on the copepod Eurytemora hirundoides, (3) Cothurnia gracilis, (4) Cothurnia havniensis, (5) Epiclintes retractilis, (6) Aegyria monostyla, (7) Euplotes harpa, (8) Aegyria oliva, (9)Amphisia pernix, (10) Stichochaeta pediculiformis, (11) Oxytricha pellionella. Suctoria: (12) Acineta tuberosa. T|ntinnoidea: (13) Parafavella elegans, (14) Parafavella edentata, (15) Ptychocylis minor, (16) Salpingella ricta, (17) T|ntinnus tubulosus, (18) Dictyocysta magna. Radiolaria: (19) Challengeron neptuni, (21) Acanthometron pellucidum, (24) Acanthochiasma fusiforme, (25) Hexalonche philosophica. Foraminifera: (20)Nonion pompilioides, (22) Globigerina bulloides. Silicoflagellata: (23) Dictyocha fibula, (26) Distephanus speculum, (27) Thalassicola nucleata. All to the same scale except (2b) and (27) (Fraser 1962) used with permission.

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Fig. 2 Small jellyfish (1) Hybocodon profiler, (2) Sarsia tubulosa, (3) Dipurena ophiogaster, (4) Podocoryne borealis, (5) Leuckartiara octona, (6) Bougainvillia principis, (7)Amphinema rugosum, (8) Eutima gracilis, (9) Eutonina indicans, (10) Aglantha digitale. (2), (3), (4), (6) and (7) to the same scale as (1); (8)^(10) to the same scale as (5) (Fraser 1962) used with permission.

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Fig. 3 Large jellyfish (1) The pearl jelly, Pelagia noctiluca, (2) Chrysaora hyoscella, (3) the big stinger or lion's mane, Cyanea, (4) the cauliflower jelly Rhizostoma octopus, (5) the moon jelly Aurelia aurita. (1) Rarely exceeds 3 inches, (5) reaches about 1 foot, but the others may occasionally be up to 3 feet across.

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Fig. 4 Other Coelenterates etc. Scale lines all represent 1 cm. Siphonophores or pseudo-siphonophores: (1) Chelophyes appendiculata, (2) Physophora hydrostatica, (3) Agalma elegans, (4) Physalia physalis, the Portuguese man o' war, (5) Velella velella, the by-the-wind sailor. Ctenophores (comb-jellies): (6) Pleurobrachia pileus, the sea gooseberry, (7) Beroe cucumis. (8) A larva of the sea- anemone Cerianthus, called Arachnactis larva. (9) A pelagic polychaete worm, Tomopteris helgolandica. The scale lines all represent 1 cm (Fraser 1962) used with permission.

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Fig. 5 Various kinds of zooplankton Harpacticoid and cylopoid copepods: (1) Microsetella norvegica, (2) T|griopus fulvus, (3) Idya furcata, (4) Oithona similis. Chaetognatha: (5) Sagitta elegans. Pteropoda (shells): (6) Clio pyramidata, (7) Spiratella (=Limacina) retroversa, (8) Spiratella helicina, (9) Clione limacina. Small Cephalopoda: (10) Sepiola, (11) Brachiotheuthis. The arrow in (4) indicates the joint between the cephalosome and abdomen

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Fig. 6 Various planktonic Crustacea etc. Cladocera: (1) Podon leuckartii, (2) Evadne nordmanni. Ostracoda: (3) Conchoecia elegans. Insecta: (4) Halobates micans. Amphipoda: (5) Themisto abyssorum, (6) Hyperia galba male (the female is much broader). Isopoda: (7) Eurydice pulchra, (8) Idothea balthica. Copepoda: (9) Caligus rapax, the sea louse, a semi-parasitic copepod. Cumacea: (10) Dyastylis rathkei (Fraser 1962) used with permission.

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Fig. 7 Calanoid copepods (1) Pseudocalanus elongatus, (2) Centropages typicus, (3) Acartia longiremis, (4) Temora longicornis, (5) Eurytemora hirundoides, (6) Calanus finmarchicus, (7) Metridia lucens, (8) Candacia armata, (9) Pareuchaeta norvegica, (10)Anomalocera patersoni male, (11) female. The arrow in (1) indicates the joint between the cephalosome and abdomen.

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Fig. 8 Crustacean larvae etc. Copepoda: (1a) and (1b) 1st and 3rd stages of the nauplius of T|griopus, (2) 4th nauplius of Calanus, (3) 5th nauplius of Centropages, (4) 5th nauplius of Oithona. Cirripedia: (5) 2nd nauplius of acorn barnacle, Balanus, (6) cypris stage of same, (7) adult goose barnacle, Lepas. Decapoda: (8) 3rd zoea of crab, Portunus puber, (9) megalopa of same, (12) phyllosome of spiny lobster, Palinurus. Euphausiacea: (10) 2nd nauplius of Thysanoessa inermis, (11) 2nd calyptopis of Thysanoessa longicaudata

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Fig. 9 Crustaceans Mysidacea: (1)Gnathophausia zoea, (2) Gasterosaccus sanctus. Amphipoda: (3)Mimonectes loverni, an exotic species. Euphausiacea: (4) Thysanoessa inermis, (5)Meganyctiphanes norvegica. Larvae of various Decapoda: (6) Sergestes, an oceanic prawn, (7) and (8) Eupagurus, hermit-crab, (9) Pontophilus, (10) Nephrops, Norway lobster, (11) Porcellana, pea-crab, (12) Crangon, shrimp, (13) Galathea, (14) Munida, squat lobster

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Fig. 10 Planktonic Tunicata Appendiculata: (1) Oikopleura dioica, (2) Fritillaria borealis. Thaliacea: (3) Thalia democratica, solitary stage showing the budding stolon (a) which gives rise to a chain of aggregate stages, (4) Thalia democratica, aggregate stage with a single embryo (b) which will eventually become a free solitary stage, (5) Salpa fusiformis, aggregate stage, (6) Salpa fusiformis, solitary stage, with stolon, (7)Dolioletta gegenbauri, gonozoid, complete with sexual organs. The eggs from this stage hatch into `tadpoles'. (8) A later stage of the `tadpole' showing the remains of the outer capsule and the oozoid developing inside, (9) the late oozoid or `old nurse' now devoid of almost all its internal organs, but with broad muscle bands and balancing organ (statocyst), a nerve center and the remains of the dorsal process, (10) a view from above of the oozoid at its functional stage with a prominent dorsal process, (a) the buds developing on the stolon, (b) the buds migrating to the dorsal process, (c) double rows of lateral zooids, the trophozoids which serve only to catch food for the whole, (d) median rows of phorozoids, and (e) the youngest migrating buds which will become new gonozoids, (11) a later stage of a phorozoid, now broken free from the dorsal process of the oozoid and acting as a bearer for two developing gonozoids. These will eventually become the free living sexual form figured in (7) (Fraser 1962) used with permission.

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Fig. 11 Planktonic larvae of various invertebrates (1) Pilidium larvae of nemertine worm, (2) Actinotrocha larva of Phoronis, (3) Cyphonautes larva of a sea moss,Membranipora. Gasteropoda (univalve mollusks): (4) Nassarius reticulata, (a) larva just after hatching, (b) old larva with shell forming, (5) Nassarius incrassata, old larva, (6) Actis minor, old larva with shell forming, (7) Littorina littorea (periwinkle), (a) young larva, (b) later larva. Lamellibranchiata (bivalve mollusks): (8) Tellina sp., (9)Macoma baltica, (10) Cardium edule (cockle), (11) Mytilus edulis (mussel), (12) Pecten opercularis (queen, or small scallop). (1)-(7) from Thorson, (8)-(12).

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Fig. 12 Planktonic stages of polychaete worms (1)Autolytus prolifer, with eggs, (2) Harmothoe imbricata, (a) metatrochophore, (b) nectochaete stage, (3) Phyllodoce groenlandica, old trochophore, (4) Phyllodoce maculata, old larva, (5) Nereis pelagica, nectochaete stage, (6) Nephthys ciliata, (a) young trochophore, (b) and (c) metatrochophores, (7) Polydora coeca, old larva, (8) Pygospio elegans, old larva, (9)Myriochele danielsseni, young larva, (10) Nephthys coeca, nectochaete stage, (11) Megalona papillicornis, old larva, (12) Pectinaria auricoma, (a) young larva, (b) old larva in a gelatinous tube, (13) Lanice conchilega, old larva in a gelatinous tube. All from Thorson (Fraser 1962) used with permission.

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Fig. 13 Echinoderm larvae etc. (1) Bipinnaria larva of a starfish, Asterias glacialis, (2) Brachiolaria larva of Asterias rubens, (3) Auricularia larva of a sea cucumber, Synapta digitata, (4) Doliolaria larva, or pupa, of Synapta, (5) Pluteus larva of a brittle star, Ophiura texturata, (6) Pluteus larva of a brittle star, Ophiothrix fragilis, (7) Pluteus larva of a sea urchin, Echinocardium cordatum, (8) Tornaria larva of the acorn-worm, Balanoglossus (Hemichordata).