A MARINE BIOLOGY

FIELD COURSE MANUAL

Extracts from a manual created for the use of first year undergraduates from the University of York whilst at the University Marine Biological

Station, Millport (Great Cumbrae in the Firth of Clyde).

by

Dr CJC Rees and Dr PJ Hogarth

CONTENTS page TIDES & WATER MARKS 3 DISTRIBUTION AND ABUNDANCE OF ROCKY SHORE ANIMALS 8 EXPOSURE 9 SHORE PROFILE RECORD SHEET 11 QUICK ESTIMATES OF ABUNDANCE OR ORGANISMS (ACFORN) 12 EXPOSURE INDEX 13 PARTICULATE SHORES 15 PHYSICAL AND CHEMICAL PROPERTIES OF SANDS AND MUDS 15 PARTICLE SIZE AND DISTRIBUTION 16 POROSITY AND WATER CONTENT 17 SOLUTES 17 TEMPERATURE 18 BEACH SLOPE AND WAVE ACTION 18 TAKING QUANTITIVE SAMPLES

OF LARGER BURROWING ORGANISMS 19 GATHERING PHYSICO-CHEMICAL MEASUREMENTS 19 DISSOLVED OXYGEN 20 BEACH PROFILE 22 LAB. ANALYSIS FOR OXYGEN CONTENT OF SEA WATER (table) 23 MEIOFAUNA 25 EXTRACTION OF MEIOFAUNA 26 PARTICLE SIZE DISTRIBUTION ANALYSIS 28 SILT AND CLAY FRACTIONS OF SEDIMENTS 29 ORGANIC CARBON CONTENT OF SANDS AND MUDS 31 DISSOLVED CARBON CONTENT 32 ESTIMATION OF SALINITY 32 Hydrometer 32 Refractometer 33 Salinometer 33 RECIPES 36 READING 38

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TIDES

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TIDES AND WATER MARKS TIDES and the EARTH-MOON SYSTEM The earth-moon system rotates in space about its common centre of mass, there being a stabilising balance of centrifugal force tending to pull it apart with the gravitational attraction between the two bodies tending to keep it together. On the side of the Earth nearer the Moon, there is a slight excess of gravitational force; on the side opposite the moon there is a slight excess of centrifugal force. These create tidal bulges in the ocean, below which the Earth is rotating about its polar axis, so that the bulges (separated by 180º of longitude) seem to travel around the Earth. The bulges (tides) are not separated by exactly 12 hours because the Moon is revolving round the Earth as well, so that the bulges are actually separated in time at any one point on the Earth's surface (in theory) by about 12 hours and 26 minutes. The Moon appears at its highest point in the sky (azimuth) over any point on the Earth's surface 52 minutes later each Earth day (Figs 1a and lb).

TIDES and the SUN-EARTH-MOON SYSTEM

The Sun can also cause tidal effects (Fig. 2a). When the Sun, Earth and Moon are all in line in space, the tidal forces are greater and the tidal bulges larger than when the Sun, Earth and Moon form a right angle in space. All three are in line at new moon and full moon, and it is then (or slightly after, for much more complex reasons) that the biggest tidal ranges (high to low water mark) are. These are SPRING TIDES (NOTHING to do with the season; from a Norse word meaning “swell” (in the English, not American, sense). New and full moon are separated by half a lunar month (27.3 days/2 = 13.65 days) so Spring tides come round approximately every 13½ days. Halfway in time between new and full moons are the first and third quarters [Half Moon]. The Sun, Earth and Moon then form a right angle in space (Fig. 2b), and the tidal forces are at their smallest, because Sun and Moon are pulling at 90° to each other. The smallest tidal ranges happen then, called NEAP TIDES (derived from another Norse word, meaning scarce). These are also separated in time by about 13½ days. TIDE LEVELS There is a MEAN TIDE LEVEL (MTL) about which the tidal oscillation is, on average, symmetrical. This is in mid-shore. High waters of SPRING and NEAP tides leave different high water marks. These are MEAN HIGH WATER SPRINGS (MHWS) and MEAN HIGH WATER NEAPS (MHWN). Similarly, with low waters, we have MEAN LOW WATER SPRINGS (MLWS) and MEAN LOW WATER NEAPS (MLWN). Particularly high or low tides occur when the Sun is at its nearest to us at the Equinoxes and create EXTREME high and low waters (EHWS or EHWN, ELWS or EHWN). Classically, these happen close to the Vernal (21 March) and Autumnal (21 September) Equinoxes. These abbreviations appear frequently in marine biological writings and on Ordnance Survey maps.

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THE EARTH, THE MOON AND THE TIDES Fig. 1a Tide-generating forces

WATER

MOON

Tidal bulge due to excess of gravitational force over centrifugal, in rotating earth-moon system.

OF COUbut the effe

EARTH

Tisudi

Axis of rotatiowhat is calledrotating system

RSE, the tidal water bulgect is the same, and at a hum

What is shown here for the Moon, alsoapplies in most respects to the effect ofthe Sun on the Earth's oceans (Fig. 2).

des are maximal at the point on the Earth'srface immediately ‘beneath the Moon', or at theametrically opposite point on the Earth's surface.

n of Earth-Moon system This runs through the barycentre (mass centre) of the whole. The barycentral axis is well inside the Earth.

Tidal bulge due to excess of centrifugal force overgravitational, in rotating earth-moon system.

Axis of Earth’s rotation

s shown above are ENORMOUSLY exaggerated, an scale tidal forces are pretty big, as are their sizes.

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Fig. 1b Why High (and Low) tides recur somewhat later (52 minutes) each Earth Day

In the diagram below, imagine that you are looking down onto the Earth-Moon system from above.

The Earth is rotating about its own axis, and the Earth-Moon system is rotating about its own axis, in the same direction in space.

3. In one earth day, the Moonmoves through about 13º of its orbitround the Earth, from M to M1.

4. During this Earth day + 52 minutes, our point will have passed through two high tide positions (One under the Moon, and one when diametrically opposite it).

This means that each high tide is separated by approximately 12 hours + 26 minutes.

M

M1

EARTH

.P1

P

13º

1. In one Earth day, a given pointP on its surface moves through360º as the Earth rotates.

2. However, the Moon has gonea further 13º in that time (see 3.opposite). So, to catch up, andagain be ‘beneath the Moon’, ourreference point on the Earth hasto go a further 13º. This takes the52 minutes by which the tidesare later each full Earth day. Thepoint has now reached P1.

[Lunar Cycle: Orbit Period about 271/3 days. So 360/27.3 degrees per day = about 13º]

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SPRING AND NEAP TIDES The Position of Earth, Sun and Moon

Fig. 2a The Position at SPRING tides.

WHEN THE SUN-EARTH-MOON SYSTEM IS IN ALIGNMENT

FULL MOON

Fig. 2b The Pos

WHE

HALMOO

LUNARTIDE

NEW MOON

EARTH SUN

This happ

ition at NEA

N THE SUN

F N

EARTH

This also ha

Summed lunar and solar tides reinforce each other tocreate a bulge. This gives rise to SPRING TIDES.

ens twice every lunar month, once every 13½ days.

P tides

-EARTH-MOON SYSTEM FORMS A RIGHT ANGLE

LUNAR TIDE

SUN

Summed lunar and solar tides detractfrom each other, so the overall tide bulge is smaller. This is a NEAP TIDE.

HALF MOON

ppens twice every lunar month, once every 13½ days.

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THE

ROCKY SHORE

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THE DISTRIBUTION AND ABUNDANCE OF ROCKY SHORE ANIMALS

Many shores, rocky or otherwise, are affected by cyclic tidal submergence and emergence. All shores are affected, more or less, by waves, which are caused by wind.

When the tide is IN -

• The temperature to which intertidal organisms are exposed is relatively constant. Physiologically high temperatures (above 30°C) are most unlikely.

• Water loss by evaporation stops (although losses by osmosis can still occur, if an organism is liable to them.)

• Availability of oxygen is determined by its concentration and diffusibility in huge volumes of sea water, likewise CO2, and it will not vary much.

• Waterborne nutrients become available.

• Light energy fluxes are reduced, and some wavelengths such as ultraviolet and infrared more than others, the extent of spectral change depending on the depth, colour and turbidity of the water.

• Buoyancy of water is available, and the swimming of larger animals is possible. Large intertidal algae can be buoyed up towards the light (hence air-bladders).

• Swimming predators can operate, swimming gametes and settlement phases can work and filter feeding is possible.

• Mechanical action of waves and currents can take place, and be liable to affect the attachment of all sorts of organisms.

When the tide is OUT -

• The temperature is liable to fluctuate more (maybe going below freezing point or above physiologically tolerable levels) . Shallow rock pools could reach 30+°C.

• Desiccation by evaporation is possible; so is evaporative cooling.

• Oxygen availability is determined by its concentration and diffusibility in air or by relatively sudden changes in small bodies of water. Possible anaerobiosis.

• Waterborne nutrients are cut off, unless proper roots exist.

• Light energy fluxes are greater, and both UV and IR irradiation may be severe. Radiative overheating is more likely.

• Buoyancy of water disappears, floppy organisms collapse and swimming (except in pools and water films) is impossible.

• Terrestrial and other air-breathing predators can operate; filter feeding has to stop, except in pools and water films. Browsing is still possible.

• Wave action ceases and sessile objects tend to stay put.

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EXPOSURE

“EXPOSURE” is a blanket concept which incorporates most or all of these factors, although it has come to be associated more with extents of wave mechanical action, desiccation and thermal/light irradiation, fluctuation and extremeness.

EXPOSURE MODIFIERS When waves can act: WIND

• Direction of the wind and the length of uninterrupted sea (the ‘fetch’) in that direction (the larger the fetch, the bigger the waves) .

• The extent to which the fetch is large in all directions from the shore (points are more exposed than bays) .

• The wind velocity and persistence from one direction (the bigger, the larger the waves) .

SHORE TOPOGRAPHY

• The slope of the shore (rate at which energy in waves is dissipated per unit area of shore surface is greatest on steep shores).

• Roughness and channelling of the shore (local current concentration).

• Algal bed wave dampers. (dense growths reduce wave action on other organisms).

TIDAL TIMING

• The time of day at which low and high spring tides are likely to occur: e.g. midday low tides make for relatively greater exposure to irradiation.

When waves cannot act: GENERAL

• All the terrestrial climatic variables (see above) including the presence of freshwater run-off or rain.

SHORE TOPOGRAPHY

• The slope of the shore (affects drainage rate and desiccation liability).

• Roughness and channelling of shore (as above: can create sheltered areas).

• Algal heaps (reduce extremity of exposure when collapsed for those things which happen to be underneath).

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All the tidally affected phenomena operate to extents which vary with position up the shore. Submersion varies between 100% and 0% of the total time. Shores experience a complex of environmental gradients. However, biological indicators are probably the best bet for trying to attach a quantitative estimate of exposure to a particular shore to enable general comparisons between sites.

Biological indicators appear in the interlinked distribution and abundance patterns for various shore organisms. Distribution patterns can manifest themselves as zonation, but of course abundance can determine whether an actual zone is recognizable or not.

When working on the shore, do keep in mind adaptation of organisms to all these influences. You can certainly do this without knowing what an organism is called, but it is important that you become familiar with as many organisms as possible: indeed, it becomes almost impossible to make sense of the marine environment without being reasonably familiar with the identities of the most significant organisms, and their key anatomical - physiological - adaptational features. Scientific names are actually an aid to remembering organisms, not an impediment!

So identification is important. Use the many field guides available (see READING p. 38). Pretty soon you will realize that there are a great many organisms that you can’t adequately identify by looking at pictures. At this point, you will need to use more serious identification keys. They can be off-putting but if you persevere most keys can be worked through effectively. Proper identification is crucial to science. There’s a limit to what you can usefully say about an organism if you don’t know what it is!

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11

12

13

SANDY and other particulate

SHORES

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PARTICULATE SHORES (sedimentary shores)

Sandy and muddy shores look devoid of life in comparison with rocky shores, but this is deceptive. The lack of large stones and exposed bedrock means that epibiotic attaching organisms such as the larger algae, barnacles, mussels and so forth, cannot survive. Sands and muds are mobile substrates offering little in the way of surface attachment sites, except where water current flow over them is permanent, very gentle and constant in direction. These conditions are met in many abyssal locations, where stalked crinoids, sponges, coelenterates and sea squirts can all attach to the sediment with forms of holdfast, but never intertidally on sandy or muddy shores.

Sands and muds are, however, penetrable and allow a burrowing INFAUNA to live there. Epifauna and epiflora are replaced by infauna. ‘Plants’ buried even a few millimetres deep in sand will receive hardly any light for photosynthesis, and will be microscopic, if present at all. Burrowing infaunal animals are usually thought of as those which are significantly larger than the sediment particles, having to force a passage through, with maximum dimensions in the range 1-10 cm, roughly. These are animals like certain POLYCHAETE ANNELID WORMS, ECHINOID ECHINODERMS, CRUSTACEA and BIVALVE MOLLUSCS.

There is also another group of infauna which are so small as to be able to move freely in the liquid filled spaces between the particles of sand. These are the MEIOFAUNA (or interstitial fauna) of NEMATODES, PROTOZOA, certain small CRUSTACEA and ROTIFERS (among others) which may be very abundant, but hardly ever noticed unless special techniques are used to reveal them. Their maximum dimensions are often between 0.05 mm and 0.5 mm (see p. 25).

Primary production may be bacterial, sometimes fungal, but seldom relying on photosynthesis. Feeding of consumers may rely on waterborne, filterable living or dead organic material from the main body of sea water, or detritus which has landed on the sand, or become incorporated in it by current action. Otherwise infaunal predators may eat infaunal prey, or they may both emerge from the sand for part of their lives to feed and/or reproduce.

PHYSICAL AND CHEMICAL PROPERTIES OF SANDS AND MUDS Sands and muds are a mixture of mineral particles separated by water or gas filled spaces. Substrata like these can be defined in terms of:

1. Mean Particle Size.

2. Particle Size Distribution.

3. Total Pore Space.

4. Water Content, and within the water.

5. The concentrations of a variety of chemical components, such as: a) Dissolved salts (NaCl, KCl etc.) b) Oxygen c) Organic carbon d) Nitrogen in various forms (may overlap with (a)) e) Sulphide f) Carbon dioxide.

6. Temperature profiles (how temperature changes with depth down into the sediment)

7. Light Penetration.

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Work on the particulate shores can involve the measurement of most of these. Tests can be done so as to provide a survey of changes in these qualities across and up and down the shore, as a function of depth within it, and if time permits, in time showing diurnal, weekly, monthly or annual changes. Physicochemical data about environmental gradients provides information against which to examine the data about animal distribution.

Factors 1 to 4 can affect:

A. How easily the sediment can be penetrated by a burrowing animal B. How easily any burrow made in it will collapse, and so whether it may need to be lined in some way C. How easily gases and nutrients etc. can diffuse through the spaces between the particles, and how easily the surface layers will dry out. D. Whether organisms will tend to be milled to pieces by the mobile grains, when they are disturbed by wave action.

PARTICLE SIZES and SIZE DISTRIBUTION

These are usually graded according to a logarithmic diameter series, based upon a 1 µm unit, called the Wentworth Scale.

Particles Diameter (µm) and sieve mesh

that retains them φ = (-log2 (mesh size in mm))

Granules

4000 2000

-2 -1

Sands

1000 500 250 125

0 +1 +2 +3

Silts

62 4

+4

+8 Clays

< 4 >+8

If all the particles were the same size, and spherical, we would have the extreme in WELL SORTED material. There would, theoretically, be about 25% of pore space between the packed spheres.

Where the particles are all sorts of different sizes, we have varying degrees of BADLY SORTED material. These tend to have smaller pore space, because the smaller particles can fit into the interstices between the larger ones.

Coarser particle sands tend, when poked mechanically, to firm up (The whitening and ‘hardening’ of some sands when they are walked on is an indication of this.) The more an animal probes at a sand such as this, the more it will find its progress resisted. These are DILATANT sands.

Finer sands, conversely, tend to become more fluid when disturbed (the quicksand effect). These are THIXOTROPIC sands and muds and will become easier to burrow into the more the animal moves about.

Animals can locally increase the thixotropy of a sediment by jiggling it, or blowing water at it; this sorts the sand, and brings finer particles to the point of disturbance. Molluscs and Crustacea make use of this quite a lot.

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POROSITY and WATER CONTENT

Porosity is the space not filled by mineral grains as a percentage of the total volume of the sample. It can be the same as the water filled volume %, but coarse, well-drained sands, gas may make up some of it. In finer sediments, the pore spaces get individually smaller, although the porosity may not, overall, be as low as in a coarse, badly sorted sand. Capillary action may ensure that, even when the tide goes out, the surface of the sand stays wet, and that individual grains do not touch each other.

In very fine sediments, water content can be very high. This is because a particle has associated with it a certain thickness of water film - a kind of macroscopic ‘hydration shell’. So exaggerating this, we compare

Porosity, inter-grain channel width and water content all affect availability of chemicals, including oxygen, and consequently the kinds of organism that may live in a particular grade of sediment.

SOLUTES This is a complex business. Nutrients, oxygen, metabolic waste products and osmotic pressure (as salinity etc.) are all involved, as well as pH, and can all be separately assessed. Two aspects are important:

A. Actual local concentration B. Rate of resupply, whilst local consumption is going on (or rate of accumulation, due to different effectivenesses of removal). This is affected particularly by rates of diffusion in the pore spaces, in a gas or in water.

It is often very hard to tell which, if any, of several chemical factors may be influencing the distribution of a particular infaunal animal. The polychaete worm Nereis diversicolor (ragworm) prefers low salinities. Arenicola marina (lugworm) is tolerant of very low oxygen concentrations in the sand around it, but is, of course, still connected to the well-oxygenated seawater in its burrow, so whether it still counts as ‘tolerant’ is less clear.

Oxygen may only be present at very low concentration compared with air or the main body of sea water. Finer sands tend to be more poorly oxygenated because of diffusion slowness in the smaller pore spaces. There may be far less oxygen a few centimetres down in the sand than there is at the surface. Higher temperatures in the sand encourage the metabolic use of oxygen, and can bring about depletion. The retreating tide, over fine sediments especially, is often associated with rapid decline in interstitial oxygen in the exposed sediment. Locally, there may be no oxygen at all.

Sulphides may be present at high concentrations. Sands and muds, especially the finer ones, can contain billions (109) of bacteria per gram of sediment. Many of them can reduce sulphate (SO4

=) to hydrogen sulphide (H2S). SO4

= is abundant in sea water, where the average concentration of is about

28 Mm L-1 (2.87 g L-1). H2S is toxic to most eukaryotes. We see evidence of the sulphides as a black or grey colour in the sand because the H2S becomes fixed as iron sulphides, especially at any depth in the sediment. Nearer to the surface, the yellow colour results from oxidised iron (e.g. Fe2O3.)

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Measurement of the depth at which the black or grey layer begins (measuring from the surface) provides a coarse indication of the oxygen status of the sediment, and is the depth at which the local redox potential changes from positive to negative.

TEMPERATURE Neither sand nor water has bad thermal conductivity, so you might expect rapid effects due, say, to solar radiative heating when the tide is out, so that the sand would heat up quickly. However, the specific heat of water is very large, and this greatly limits the size of the temperature fluctuations, at least on the time scale of the period between tidal inundations. Sands and muds present much less stress to their infauna in this respect than epifaunal animals may experience on a rocky shore. The extreme surface layer of a well drained sand may get too hot for anything to live there, if it is sunny whilst the tide is out, but this really only applies to the top few millimetres.

Temperature, and the way it changes with depth in the sediment are easily measured with an electric thermistor-probe thermometer.

BEACH SLOPE AND WAVE ACTION STEEPER beaches are exposed to GREATER wave action and their particles tend to be coarser. Whilst this topic is extensive, the following summary can be made:

STEEPER SLOPE AND MUCH WAVE ACTION

ATTRIBUTE SHALLOWER SLOPE AND LITTLE WAVE ACTION

coarser Mean particle size finer

higher Permeability lower

weaker Capillary action stronger

tends to drain Water content tends to stay wet

more Oxygen less

few Bacteria lots

little Sulphides much

absent, or deep Black/grey layer present at or near top

little Organic material much

A TYPICAL SANDY SHORE EXERCISE The aims are:

a) To obtain quantitative samples of the larger infaunal organisms from a series of sampling stations regularly spaced along a line transect up the shore slope. These will then be used for counting identified animals in the laboratory. The results form the basis for plots of animal distribution for comparison with any environmental gradients which have also been measured, and may reveal the presence of zonation of animals. The samples could also be used for simple estimations of biomass per unit volume of sand.

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b) To find the shore slope profile along the transect line by means of a simple levelling procedure, much as may be done on the rocky shore. This allows us to estimate the length of time for which any sampling station in uncovered by the sea (mean daily percentage).

c) To measure the values of a number of physical and chemical parameters at each sampling station. These allow coarse environmental gradients to be plotted for comparison with the distributions of animals.

1. TAKING QUANTITATIVE SAMPLES OF LARGER BURROWING ORGANISMS TYPICAL EQUIPMENT: Spade, 2 mm sieve, 1 mm sieve, white tray, a paint brush, a marker pen and some large polythene bags. The bags are best labelled in advance with sampling station, sieve size and number of samples, to minimize error and the necessity to write on wet bags.

The transect is measured out and a labelled bamboo pole put in at each sampling station. The distance between stations is decided after consideration of shore slope, time available and desired sampling accuracy. Sample up the line of canes, carrying out the same sampling procedures at (or near) all canes, even though there may, at some, seem to be few animals present. These low numbers are important in recognising any zonation pattern when the field samples sorted out in the lab.

Dig out enough sand to fill a sieve completely, and level it off. Please try not to dig just from near the sand surface. The intention is to fill the sieve with a block of sand as similar to the shape of the sieve as possible (e.g. 30 x 30 x 10 cm). Sieve out the sample by rocking and jiggling the sieve up and down in water (either in the sea or in a pit which can be dug to collect water locally. You may prefer to do this if you are far from the water’s edge but using the sea itself is easier). Remove any large stones, wipe off any loose sand around the rim of the sieve and then invert it over the white tray. Bang the bottom of the sieve several times to dislodge the rest of its contents (including all the animals) into the tray. Check over the sieve for any (especially worms) that failed to come out, extract them by hand and add them to the white tray. Never try to remove animals from a sieve with the paint brush as it fragments them and mashes them in the mesh.

Repeat the process until you have a prescribed sample in the tray. Next wash, tip or brush (now you may use it) the combined 2 mm sample or combined 1 mm sample into its correct, pre-labelled polythene bag. It is best if some sea water goes in as well, but not too much, which will make catching animals difficult later. Close the bag by tying its mouth in a knot. Use a loose-ish loop knot: remember it has to be opened again.

Return all bags to the laboratory for analysis.

2. GATHERING PHYSICO-CHEMICAL MEASUREMENTS TYPICAL EQUIPMENT: Spade, plastic 30 cm rule, two 50 ml sampling syringes, each fitted with a long sampling tube instead of a needle, a 100 ml measuring cylinder, an hydrometer (graduated from 1·000 to 1·050 g ml-1, a pH meter, a pNa meter (if available), a Redox potential or oxygen electrode meter (if either is available and working), the Winkler field kit (containing 20 sample bottles, reagents and syringes) for fixing samples of water - used later in the lab to determine their dissolved oxygen content, and a thermistor probe thermometer (or alcohol-thermometer). All these devices can be kept together, protected from breakage, sand, sea water etc. in a large plastic tray or similar, fitted with a rope so that it can be dragged over the sand. Make sure that one member of the team has a clipboard, pen(cil) and a data sheet.

(a) Dig a hole at the same distance down shore as the cane; you don’t need to do this exactly where the cane is. Avoid animal-gatherer’s diggings! Wait for water to collect in the hole. At a few stations very high up the shore this may take some time to happen. When it does:

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(i) Fill up the measuring cylinder to the top, using one of the two 50 ml syringes. Insert the hydrometer, and record the specific gravity (density) of the water. This allows you to find its salinity (for all ions combined) easily.

(ii) Measure the pH of the water, using the meter. Each time you do this, rinse off the electrode with distilled water. If there is a pNa meter, do the same thing with it.

(iii) Fix a sample of the water for determination of the dissolved oxygen concentration by Winkler titration. This procedure is the most awkward of the measurements you need to make.

(iv) Measure the temperature at depths of about 10 cm and about 3 cm below the surface with the thermistor probes or thermometer.

(b) Cut as clean a profile of the sand as you are able, and measure the depth from the surface at which a black or grey layer begins. If you cannot see one within 30 cm of the surface record that as > 30 cm. The hole will fall in too easily if you dig deeper.

(c) With such a hole, GENTLY push the Redox potential meter in between the deepest sand grains, but NOT immersing it in the water by more than 2 cm. Read off the redox potential on its screen, it being essential also to record whether this was +ve or -ve. Wash off the electrode of the redox meter with distilled water after each use.

(d) You should return to the laboratory with as many bottles labelled with their station number as there were biological sampling stations, and the data sheet with all other possible measurements. (Measurement of oxygen concentration, in ml L-1, is done in the lab. All you collect in the field are the chemically-fixed samples.)

Ideally, you will have collected data about salinity, pH, temperature, depth of anoxic sand, pNa and sand redox potentials representing environmental conditions at each biological sampling station.

MEASURING THE DISSOLVED OXYGEN CONTENT OF WATER The Winkler method for finding out dissolved oxygen concentration is quite reliable (see RECIPES p. 36). In this, iodine is released so that one molecule of iodine (I2) is equivalent to one atom of oxygen (O) originally present. The iodine is titrated against sodium thiosulphate, with a starch indicator.

Samples of sea water collected in the field have their oxygen content ‘fixed on the spot’ as iodine equivalent, for measurement later in the lab. If not, the concentration of oxygen in a sample is prone to change quickly (usually downwards) inside a sealed bottle.

TAKING SAMPLES This is done by using a 50 ml plastic syringe with a long nozzle (a plastic tube) which is freed from air bubbles by ejecting them from the syringe held point up. An important matter is that you should avoid allowing oxygen from the air to contaminate the sample. Some of your samples come from oxygen-deficient water (maybe anaerobic, even) and will tend to pick up oxygen easily, even from stray air bubbles.

Having taken your 50 ml, run this gently into a glass Winkler sample bottle to fill it completely, including the neck. After this, replace the stopper. (You can’t leave it like this, even though sealed, as any bacteria in the water could go on using up the O2 or any green algae could go on and make more.) So, the oxygen has to be chemically ‘fixed’, here in the analogous form of iodine using syringes fitted with long, square-ended, blunt needles.

Unstopper the bottle and add 0.7 ml of MnCl2 from its 1 ml syringe and then 1.5 ml of alkaline KI solution from its syringe, so that the solutions go well down into the sample bottle. Replace the stopper so that NO air is trapped. Some fluid may come out. Never mind. Roll the bottle

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horizontally between your hands to mix the beige precipitate with the water. This precipitate absorbs all the oxygen. Leave it to do this for about six minutes.

Next, carefully remove the stopper. Add 2 ml of phosphoric acid from its syringe, again getting it well down into the bottle. Replace the stopper securely and shake the bottle until the acid has dissolved the precipitate. It dissolves the iodine in so doing and your yellow sample is now safe from any aerial contamination at all, even if opened. Label the bottle with its station number and make sure the stopper is firmly in place. The rest of the determination of oxygen concentration is completed in the lab.

Completing measurements of oxygen concentration in the lab. 1. Set up a small (25 ml if available) burette in its stand and a ‘contrast tile’ on the base of the stand. Find the bottles (W5 - Recipes p. 36) of sodium thiosulphate solution and (W4) of starch indicator and have a Pasteur pipette available for adding drops of indicator.

2. Fill the burette with M/80 sodium thiosulphate solution and run a little out to fill the tap and nozzle.

3. Have your sample bottles containing released-iodine solution ready. Label their stoppers with the same number as on the bottle itself. This is necessary for §5 (below).

4. Tip the contents of a sample bottle into a 100 ml conical flask. Rinse out the bottle with some water, to get all the iodine solution out, into the flask. Add a drop or two of starch indicator; the contents of the flask will turn blue-black. Titrate against the sodium thiosulphate until the blue colour disappears. Do this carefully, i.e. slowly and shake the flask; not many ml of thiosulphate will be needed (between 0 and 4). Don’t flood it in!

5. Find the volume of each stoppered sample bottle. Dry them all, open, in an oven. Remove and allow them to cool. Refit the stoppers and weigh. Next refill the bottles with fresh water, re-stopper, blot dry and re-weigh. Find volume by difference in mass due to the water, and assuming the density of fresh water to be 1.0 g ml-1. (Use table below for values.)

Also, see the graph of oxygen saturability of sea water at different temperatures (above).

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LAB. ANALYSIS FOR OXYGEN CONTENT OF SEAWATER, [O2]

Stoppered sample bottle (masses, in grams)

Burette levels given in ml.

Volume (ml) of thiosulphate used

Concentration of oxygen in the water

Station number

Mass (dry)

Mass (full of water)

Volume X (ml)

Start level (V1)

End level (V2)

A = (V1 - V2 )/X [O2] = (70 x A)/X (ml. / litre)

1

2

3

4

5

6

7

8

9

10

11

12

13

14

15

16

17

18

19

20

3. BEACH PROFILE MEASUREMENTS This work needs three people. Sampling stations will have been paced out at regular intervals, indicated with a marker pole. The aim is to find out what the height differences are between the stations: the distances along the sand between station poles also need to be measured.

The surveying technique is identical to that used to estimate the profiles of the rocky shores with a pair of levelling poles. One is of fixed length and used as the upper pole. The other is of adjustable length by means of a graduated wooden rod and is used as the lower pole. You will also need a 50 metre tape (or longer, if I can get one), two metal pegs (one to fix the top tape end down, the other to mark where the other end of the tape reaches), a copy of the data-sheet (below) on a clipboard, and a pen(cil).

Start with the pair of bamboos which are farthest up the shore (Nos. 1 and 2). Peg the zero tape-end down to the sand at sampling station bamboo 1, and pull out the tape down shore in parallel with the line of bamboos, close enough to them to record their numbers. Leave the empty tape-reel on the sand, wherever it gets to, and push the other peg in there.

22

(i) At each interval between sampling station bamboos, take the levelling pole of adjustable length to the lower bamboo and hold it vertically. Do the same thing with the fixed length pole at the upper bamboo. Hold the poles so that their crossbars are parallel with the sea horizon, not so that they are ‘pointing’ at it. Whoever is with the upper pole sights over the crossbar and asks the holder of the lower pole to adjust its length until both crossbars and the sea horizon are all in line.

At this point whoever has the clipboard on the data sheet records:

(a) the numbers on the flags attached to the bamboo canes (already entered)

(b) the distance along the ground tape by the lower bamboo cane (to the nearest cm) and (c) the length of wooden rod projection from the lower pole, with its sign (+) or (_).

(ii) Move down shore one interval between bamboos (e.g. from 1 - 2 to 2 - 3) and repeat what you did at the previous interval.

From bamboo

Cumulative distance along

Length of lower pole projection

From bamboo

Cumulative distance along

Length of lower pole projection

cane Nos the sand (m) +/_ (mm) cane Nos the sand (m) +/_ (mm)1 0.00 + 0 10 to 11

1 to 2 11 to 122 to 3 12 to 133 to 4 13 to 144 to 5 14 to 155 to 6 15 to 166 to 7 16 to 177 to 8 17 to 188 to 9 18 to 199 to 10 19 to 20

(iii) When you reach the interval which includes the empty reel frame, do the following. It needs two of you to work together.

Leave the lower metal peg ‘X’ where it is. Suppose the ground-tape is L metres long. One person goes to pull out the upper peg holding down the zero end of the tape, and pocket it. Each picks up either the top or the downshore ends of the tape and together walk at similar pace until the zero end reaches peg ‘X’. Fix the zero end of the tape to the sand with this peg. Take the pocketed peg down to where the empty tape reel now is. Push it into the sand just by it, having made sure that the ground-tape is fully pulled out straight.

When making further measurements of distance downshore, add L metres to each reading on the ground-tape for the canes’ cumulative distance along the sand in the table. When you need to move the whole tape down shore again, add 2 x L metres, etc.

(iv) Over the length of transect, this move will need to be made several times. Each time, remember to add a further L metres to the recorded cumulative distances of each bamboo from the first one.

(v) In the lab, plot the data, and add it to the data set for analysis. You will need to supply data both about cumulative distance down shore of the bamboo poles and the difference in height between stations.

23

4. OTHER LAB WORK ON THE SANDY SHORE SAMPLING. Identify and count, all the animals which were collected in the sieve samples. There are numerous field guides and keys to use in the identification process. Crustacea and Annelids (of which there is usually a frightening diversity) are the main areas of difficulty. Molluscs present fewer problems, as you are likely to find only a few species. First, collect and sort the animals into recognisably distinct groups, tackle the identification process and re-sort. Count and record.

SORTING AND COUNTING THE SMALLER ANIMALS, FIRST IDENTIFYING THEM USING THESE KEYS TO CRUSTCEANS AND WORMS 1. With sea water, using as little as possible, carefully wash the entire contents of the bag you

are to sort into a large plastic tray. If the sample is smaller than average, a smaller tray may be more useful.

2. Remove stones etc. to improve visibility during searching. 3. Ensure that you have a shallow, but continuous layer of seawater over the sand etc. so that

you can poke about and see animals moving. Too deep and they will be difficult to catch. 4. Shake the tray, rake the sand, anything to make every animal show itself by moving. 5. Pick all of the animals up with a wide-mouthed plastic pipette (cut the end off to suit the size

of your prey), spoon or forceps and segregate them roughly by “type” in divided petri dishes.

6. Refine your rough identifications using identification guides and keys, and sort again.

USE A TOP LIT DISSECTING MICROSCOPE TRANSMITTED LIGHT IS ONLY SUITABLE FOR MINUTE OR TRANSPARENT ANIMALS

7. If the animals move about too rapidly in the petri dish, they will eventually slow down as

their oxygen supply diminishes. If this takes too long, slow them down with a drop or two of ethanol. [Not in the tray! There you need them to be alive and moving so that you can see where they are. Otherwise, they hide among the sand grains.]

8. Count and record numbers of each species.

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MEIOFAUNA

Most of the animals we find on the seashore are at least moderately large; even the smaller Crustacea are several millimetres long. These are macrobenthos and rely on either (a) emerging from the sand as a whole when the tide covers them or (b) setting up tubular connections through the sand to the overlying water, as do the worms such as Arenicola and bivalve molluscs such as Angulus.

Sand also contains many smaller organisms, small enough to live permanently between the sand grains as an interstitial fauna, otherwise called the ‘meiofauna’. They may make up 25% of the total biomass. They are bigger than the interstitial microbenthos, which are mainly bacteria and fungi. Meiofaunal movement through the sand is thought not to displace sand particles much, if at all, and these animals tend to be within the length range 0.05 to 3.0 mm. Several phyla are present, including Protozoa, Coelenterata, Annelida, Mollusca, Arthropoda, Nematoda, Bryozoa and Gastrotricha. Most of these are elongated and rather worm like, to penetrate spaces between sand grains, even though the phyletic shape is typically seldom like this. There are a few broad, flat animals, and contractility of a body, well equipped with adhesive organs, is also common. They may move by ciliary action, as in the Platyhelminth Turbellaria, Rotifers, Gastrotrichs, Annelids and Polychaete worms, but some simply writhe about, such as harpacticoid copepods and nematodes.

What types of feeding activity are there? A few are predators, such as the small Coelenterate hydra Halammohydra. Nematodes, Rotifers, Annelids and harpacticoid copepod crustacea feed on detritus or diatoms.

Meiofaunal animals are small, and are made up of rather few cells. They produce few gametes, perhaps between 1 and 10 eggs each. Viviparity with parental care is widespread. The young are not released to follow a planktonic pelagic phase. How meiofauna are distributed remains uncertain, and they may rely upon very slow lateral spreading within the sand. Long distance transfer is not understood. The use of spermatophores, rather than general sperm release into the water is usual, presumably to avoid wastage from an animal with small size and limited gamete productivity.

PHYLUM (Genus)

1. NEMATODA (Plectus)

2. TURBELLARIA (Coelogynopora)

3. GASTROTRICHA (Urodasys)

4. CRUSTACEA Copepoda (Cylindropsyllis, a harpacticoid)

5. CRUSTACEA Isopoda (Microjaera)

6. TARDIGRADA (Batillipes)

7. BRYOZOA (Monobryozoon)

8. MOLLUSCA (Pseudovermis)

9. ANNELIDA Archiannelida (Nerillidium)

10. COELENTERATA (Halammohydra)

11. ANNELIDA Polychaeta (Psammodrilus).

25

EXTRACTION OF MEIOFAUNA (interstitial animals) Meiofauna can be removed alive, but the technique is more awkward than collecting them dead and stained.

1. Dead, stained organisms (see RECIPES p. 36)

Put about 2 cm depth of wet sand sample in a 250 ml conical flask, and add 50 ml of 0.1% rose bengal solution in 5% seawater-formalin. Shake to mix thoroughly, and leave for about 2 hours. This kills and stains the meiofauna.

Decant the fluid off through a 64 µm sieve (or similar, as available), but avoid getting any sand in it. Add a further 100 ml of seawater to the flask, shake up again and allow to settle. Then decant again through the sieve. Repeat this procedure about five times.

You should now have a concentrate of meiofaunal animals, stained pink, in the sieve. To get them off, place the sieve in a petri dish of seawater, jiggle it to and fro and suck out the animals which float off with a Pasteur pipette into a suitable small tube. Examine them in a cavity slide at a magnification of about x20 or x50. The sieves can be made conical to help with this, or you can quite easily use a fine plastic tea strainer instead of a 64 µm sieve plate. Animals can be lifted out with a fine paint brush.

2. Live Organisms (Considerably more awkward)

Treat about 20 g of sand with 100 ml of 3.5% magnesium chloride solution. This is isotonic with seawater and anaesthetises the animals to make them let go of the sand grains. Leave for about 10 minutes.

The extraction is now carried out either as in the method for dead animals or by serial washings and decantation through a fine sieve; you may be able to stain them, without killing them, in methylene blue, but its staining ability is highly unpredictable, especially with living Arthropods.

References : Eltringham: Life in Mud and Sand. pp. 48-54 Brafield: Life in Sandy Shores. pp. 13-15. Holme and McIntyre: International Biological Program Handbook. 16 pp. 163-168. McIntyre: Ecology of Marine Benthos. Biol.Rev. 44:245-290.

26

SOME

PHYSICAL MEASUREMENTS

27

PARTICLE SIZE DISTRIBUTION ANALYSIS This involves the measurement of the weights of fractions of the original sample of sand which are retained by variously sized sieve meshes. It also gives the water fraction of the sample. Two methods can be used: wet, ‘puddle’ sieving or dry sieving. Endecotts sieves are recommended.

To begin with, determine the wet weight of your sample, by any sensible means (weigh by difference). Then tip it out onto a piece of polythene.

For dry sieving, you must now thoroughly bone-dry the sample in an oven at a temperature of about 100°C. This involves turning over the sample from time to time with a spatula or something similar, so that all the grains become dry and separate. You then shake them through the coarsest available sieve onto a large dry sheet of paper. Retain the sample from that sieve separately. Then shake the material which passed through it through the next finest sieve, and retain that material separately. Go on like this, until you have passed the material through all the sieves available, and obtained a set of dry separately classified grain size samples. These can all be weighed separately, values entered on the chart overleaf and the sample size distribution analysed.

For wet sieving, you take the wet sample (after weighing it entire) and puddle it through the sieves in turn, retaining the graded size samples in each successive sieve. You will need a large, water filled tray to receive what falls through in each case. This then has to be poured through the next sieve, and the awkward bit is to get all of it into the sieve, rather than leaving it in the bottom of the tray. You can flush it out with a wash bottle. All the loaded sieves then need drying off, their contents tipped out, and treated as for dry sieving. Wet sieving is more thorough than dry. You get more through that would be mechanically held back by other particles in dry sieving, but it is messier and takes longer.

The values of φ corresponding to various sizes of sieve mesh are given below. φ = -log2(mesh size in mm) See Wentworth scale of particle grading earlier on in these notes.

Table of meshes and φ values A guide based on sieve sizes that might be available

Mesh (µm) φ Mesh (µm) φ 2000 -1 300 1.74 1000 0 250 2 710 0.49 150 2.74 600 0.74 65 3.94

SIEVE ANALYSIS

W E I G H T S Cumulative Sieve Mesh φ Sieve + Sieve Grains %Total % of Total

'M' (µm) Grains Dry Wt Grain Wt TOTAL:

φ values for other meshes can becalculated from the expression:

φ = -log2(mesh in µm)

28

The scale of φis standard inthis sort ofanalysis andprovides alogarithmic scale of grainsizes.

The median particle diameter, Mdφ is the φ value corresponding to 50% cumulative frequency. The spread of particle size sorting is assessed from the number of φ units lying between the 25% and 75% cumulative frequency points on the distribution curve.

25% point, Q1φ = ___________________________ 75% point, Q3φ = ___________________________

Sorting is then given by QDφ = Q3φ - Q1φ = ___________________________ 2 Small values of QDφ typify well-sorted sands. These also tend to have larger porosities. This analysis loses all particles which pass through the finest sieve. TO MEASURE POROSITY Weigh a suitable dish empty. Put in your sample and reweigh to get its wet weight by difference. (Use about 70 g) Spread out the sample and put it somewhere to dry for about 4 hours. Reweigh when cool to give the dry weight of silica, WS. Let the total wet weight be Wt.

Calculation:

"Sand" contains salt water and silica (at simplest). Take salt water density as 1.026 g ml-1 and silica density as 2.65 g ml-1.

Let the weight of salt water = WW with a volume VW ml

Let the weight of silica = WS with a volume VS ml

So: Wt = (1.026 VW) + WS and Wt - WS = VW and WS = VS

Porosity = Water filled volume = VW / (VW + VS) Total volume

So, combining measured quantities, Porosity = (Wt - WS) . 100

(Wt - WS) WS

1.206 2.65

2.65 1.206 + %

1.206

SILT AND CLAY FRACTIONS OF SEDIMENTS The particles between 62 and 4 µm in diameter are defined as SILT. Those smaller than this are CLAY. The ease with which diffusion is possible between grains (oxygen, nutrients etc.) is much affected by the amount of, silt and clay present. Clayey sediments are often very poor in oxygen, because the pore sizes (but not necessarily the porosity) are so small.

The effective limit of sieves is at about 64 µm, but you can make use of settlement rates to determine how much clay and silt there are. If you shake up a lot of fine particles in water and leave it to stand, the larger ones will settle out faster. Differently sized particles will sink at different rates.

For silica spheres: (in what follows, assumed sphericality is a source of error) 29

Particles settle through in

Diameter (µm) Distance (cm) Time 62 10 39 seconds - 32 10 116 seconds 1 min 56 sec 16 10 464 seconds 7 min 44 sec 8 10 31 minutes - 4 10 123 minutes 2 h 3 min

Thus, after 464 seconds there will be no particles larger than 16 µm diameter in the top 10 cm of suspension. However, all the other particles will be present in their original proportions, at a depth of 10 cm.

So, if we take a sample of the suspension from 10 cm down (i) immediately after shaking and (ii) 464 seconds later, then sample (i) will contain all particles and sample (ii) will contain only particles less than 16 µm in diameter. By filtering, drying and weighing we can arrive at the relative amounts of 62-16 µm particles and less than 16 µm particles in the sediment. By knowing the original weight of sediment which was shaken up, and the proportion of the total suspension which was taken in the sample, we can calculate the total weights of particles larger or smaller than 16 µm. Similar weighings after 31 minutes and 123 minutes will give estimates of the amounts of 8 and 4 µm particles present.

METHOD

1. Weigh a sediment sample; perhaps about 300 g will do. Oven dry it thoroughly, and reweigh.

2. Transfer the dry sediment to the surface of a 64 µm sieve, placed in a flat bottom basin. Add about 400 ml of water, to flood the sieve. You must not use more than one litre! Wet sieve the sediment by puddling and agitation until the fine fraction has passed through. This takes time.

3. Transfer the material in the BASIN to a stopperable 1 L measuring cylinder. Wash out any sludge into the cylinder from the basin and make up the total volume in the cylinder to 1 L with distilled water. Stand the whole thing in the lab. to reach ambient temperature for an hour (temperature affects settling rate a lot).

4. Stopper the cylinder and shake it up well. IMMEDIATELY take a 20 ml pipette sample from a depth of 20 cm. Transfer the pipette contents to a small weighed dish. Dry it out at 100°C; you can boil off the water, but do this with care.

The weight in here represents the total amount of sediment less than 64 µm size in a 20 ml sample, i.e. 1/50th of the whole lot.

5. Shake the cylinder again and leave it to stand for 7 minutes 44 seconds. Then take a second 20 ml sample from a depth of 10 cm below the surface. Transfer this to a weighed dish; dry out. (The silt fraction less than 16 µm in diameter).

6. Shake the cylinder again and take a third 20 ml sample from 10 cm depth after 2 hours and 3 minutes. Dry it off as before. (The fraction less than 4 µm).

7. If the pipette samples are 20 ml, then since the whole suspension is 1000 ml, we multiply the weights of the various extracted samples by 50.

Let the dry weight of the 1st sample = W1 (less than 64 µm) Let the dry weight of the 2nd sample = W2 (less than 16 µm) Let the dry weight of the 3rd sample = W3 (less than 4 µm)

30

Then, in the WHOLE SAMPLE:

The weight between 0 and 4 µm = 50 . W3 (CLAY) The weight between 4 and 6 µm = 50 . (W2 - W3) (FINE SILT)

The weight between 16 and 64 µm = 50 . (W1 - W2) (COARSE SILT)

These can be related to the initial sample wet and dry weights as required. If the sample is one which you have analysed with sieves, this information can be included in the cumulative particle size graph, with points for φ = +4, +6 and +8, by using your loaf a little.

It is also a simple way of doing something rather tricky otherwise, but assumes that the particles are made of quartz and that they are spherical. ORGANIC CARBON CONTENT OF SANDS AND MUDS Weigh out about 5 g of sediment, and record this as mass W (g).

Transfer it to a 500 ml conical flask. Add 10 ml of 1M K2Cr2O7 solution and then 20 ml of conc. H2SO4. Shake carefully for a minute and then put in a boiling water bath for 15 minutes. This oxidises the organic matter, and in so doing, reduces some of the dichromate.

Cool, add 200 ml of distilled water, 10 ml of glacial phosphoric acid and 1 ml of diphenylamine indicator.

Put 1M FeSO4 solution in a burette and titrate the solution in the flask slowly. The solution will go purple or blue, and the end point occurs when the solution changes to green. When this happens, add another 0.5 ml of 1M dichromate solution to restore an excess. Complete the titration by adding FeSO4 dropwise from the burette until all the blue colour disappears. PRINCIPLE

Organic matter is digested with a very acidic chromate solution. A known amount of chromate goes in at the beginning, and any which is not reduced by the organic matter is titrated against a standard ferrous sulphate solution. CALCULATION

1 ml of dichromate is equivalent to 3 mg of carbon. So, the amount of organic carbon in the sediment sample is (V1 - V2) x 0.003

W

where V1 is the volume of 1M K2Cr2O7 used (10.5 ml); V2 is the volume of FeSO4 needed; W is the mass of the sampl

Certain substances, such as coal dust, cause errors here, but the method is as sim

AN ALTERNATIVE, MORE APPROXIMATE METHOD N.B. What follows is given very much in note form. Weigh out about 50 g of sthe sample out on a clean tray and leave it to dry in a heated oven (80°C). Rewsample up again in distilled water (200 ml), pour off the water + dissolvedReweigh. Finally heat, the sample to red heat for 15 minutes, allow to cool, anConsecutive differences between original wet weight and the three re-weigfollowing:

1. Water content 2. Soluble material content 3. Insoluble organi

31

x 100%

e (g)

ple as possible.

ediment, wet. Spread eigh. Now shake the

salts and dry again. d reweigh yet again. hings give you the

c content.

DISSOLVED CARBON DIOXIDE CONTENT Carbon dioxide and water stabilise within a set of equilibria as below:

CO2 + H2O l H2CO3 l H+ + HCO3- l 2H+ + CO3

= In sea water at about pH 8, no CO2 exists freely in solution, but has become combined within bicarbonate ions, HCO3

-. Isolated volumes of sea water may accumulate carbon dioxide, to reduce the pH to 5 or 6, as may happen in rock pools.

Samples

In a sample, let the carbon dioxide be present as HCO3-. This can be reacted with an acid,

providing H+, such as hydrochloric acid. Thus:

HCO3- + H+ d H2O + CO2 at about pH 4.5

In a titration with HCl, we use an indicator passing its colour change point at pH 4.5, such as with Bromocresol Green (Yellow at pH 3.6; Blue at pH 5.2).

Size of sea sample : S ml Volume of acid solution needed to reach pH 4.5 : V ml Molarity of acid solution : M molar

Concentration of Carbon Dioxide = (V . M)/S moles L-1

1 mole of CO2 weighs 44 g, equivalent to 22.4 L at N.T.P.

Hence you can find the CO2 content in mg L-1 or ml L-1 by conversion of the concentration figure.

Solutions Required

Bromocresol Green Indicator 0.1 M HCl (dilute to make 0.01M HCl)

ESTIMATION OF SALINITY Hydrometer method A simple, inexpensive and slightly inaccurate method, making use of the relationship between the specific gravity of the water and the concentration of dissolved salts. Hydrometers are fragile glass instruments and their readings are sensitive to temperature, BUT they can be used easily in the field, and quickly, to give quite a fair estimate of salinity. You need a 1.000 to 1.050 g ml-1 hydrometer, a measuring cylinder to hold the water sample and to float the hydrometer in, and a thermometer. Hydrometers are weighted so that they float with the long thin stem vertical, and you read the specific gravity (density) of the seawater off along the graduations marked on the stem. Hydrometers break easily.

The table below allows correction to be made for temperature, salinity being given in parts per thousand (‰) of dissolved solutes, or chlorinity (g Cl L-1)

This method won't tell you which solutes (and how much of each) are responsible for the increase in specific gravity (density) of sea water above 1.000 g ml-1.

32

Graphs showing the relationship between the specific gravity of sea water at selected temperatures, and salinity and chlorinity

Refractometer method Salt water refracts light differently from pure water, and the higher the salt concentration, the bigger the difference is. Refractometers are portable, about the size of half a pair of small binoculars. They are also expensive, and easily damaged by abrasive grains of sand, or being dropped.

They have what can best be described as a hinged lid at one end, over a glass plate (End A) and an eyepiece at the other (End B).

First, put a couple of drops of distilled water on the plate, using a Pasteur pipette or its plastic equivalent, and close the lid at End A.

Next, point End A at daylight, and look through End B, keeping the closed lid on the upper side of the tube (otherwise the sample will fall out). When you look through the eyepiece you will see a darker (grey) straight edged area, this edge being in line with the scale of salinity that is also visible. It should read l.000, or 0‰, being distilled water. It can be calibrated, if not.

Next open the lid and dry off the water with soft tissue. Next, put drops of the water of which you wish to measure the salinity onto the plate at End A and close the lid again. Repeat the procedure used for pure water, and read off the salinity of the sample.

DO NOT DROP A REFRACTOMETER, GET SAND IN IT or forget to DRY OFF THE GLASS PLATE AFTER MEASURING. You will not need to calibrate it against distilled water each time.

Salinometer method Salinometers are simple to use, and rather like a portable pH meter. HOWEVER, they can be ruined instantaneously if dropped into sea water (just like a digital camera! Merryweather pers. comm.), rather than just immersing the glass bulb end.

33

Average Composition of Sea Water (Salinity 34.33‰, Chlorinity 19‰)

ION gm kg-1 Sea water mM kg-1 Sea water Na+ 10.556 459.02 K+ 0.380 9.72

Mg+ 1.272 52.30 Ca++ 0.400 9.98

Cl- 18.980 535.30 SO4-- 2.649 27.57

HCO3- 0.140 2.29

Br- 0.065 0.81 (Nitrogen) 10-6 to 0.0007 (depends on the form present)

(Phosphorus) 10-7 to 10-4 (depends on the form present)

34

These make up 99.7% of the total sea salts

RECIPES

35

SOME ADDITIONAL RECIPES WINKLER O2 DETERMINATION KIT a) 10 x 1 ml syringes 10 x 2 ml syringes 10 x 10 ml syringes 4 x 50 ml syringes 34 x #19 x 2” needles

b) W1 500 ml MnCl2 60 g L-1

47.14 g Magnesium Chloride 4H2O made up to 500ml with dist. water. Label: “W1 Winkler Manganese Chloride soln."

c) W2 500 ml NaOH/KI 128 g NaOH + 80 g KI made up together to 500 ml with dist.water. Label: “W2 Winkler Alkaline KI - CORROSIVE”

d) W3 1L 50% orthophosphoric acid 500 ml glacial phosphoric acid + 500 ml dist. water Label: “W3 Winkler Phosphoric Acid”

e) W4 1 L 10 g L-1 soluble starch in saturated salt solution Dissolve 1 g soluble starch in 100 ml saturated NaCl soln. Label: W4 Winkler Starch Indicator

f) W5 2 L 0.0125 M (M/80) Sodium Thiosulphate soln. 6.204 g Sodium Thiosulphate 5H2O made up to 2 L with dist. water or use sealed ampoule from stores to make M/80 soln. by dilution Label: W5 Winkler Sodium Thiosulphate

g) 50 x 30 ml stoppered bottles

h) 6 x 100 ml plastic bottles

MEIOFAUNA EXTRACTION: M1 Rose Bengal 1g in 140 ml conc. formalin + 860 ml sea water Label: “M1 0.1% Rose Bengal in formalin/sea water”

M2 150.5 g MgCl2 dissolved in 2 L sea water label: “Magnesium Chloride soln. (equiv. 35‰ salinity)

M3 500 g ZnCl2 dissolved in 1 L dist. water label: “Zinc Chloride soln. (50%) for floatation - POISON”

36

READING

37

BIBLIOGRAPHY identification guides - These are all useful for identifying marine species. The guides published by Hamlyn (Campbell) and Collins (Hayward et al.), and Gibson et al. are essentially field guides, relying on pictures, and including most of the common species that can be fairly easily identified. But rarer and more difficult species (of which you will find some!) tend not to be included or receive superficial treatment, and cannot be satisfactorily identified from pictures. For these, you should use proper diagnostic keys, such as are provided in the most extensive book for marine animal identification, Hayward and Ryland, 1995 (2 vol. hardback or paperback condensed version).

Campbell, A.C. The Hamlyn Guide to the Seashore and Shallow Seas of Britain and Europe. (Hamlyn: various editions).

Crothers, J. (1997). A key to the major groups of British marine invertebrates. AIDGAP Field Studies Council.

Crothers, J. & M. (1983). A key to the crabs and crab-like animals of British inshore waters. AIDGAP Field Studies Council.

Fish, J.D. and Fish, S. (1989). A Student’s Guide to the Seashore. (Unwin).

Gibson, R., Hextall, B. and Rogers, A. (2001). Photographic Guide to the Sea & Shore Life of Britain and North-west Europe. (Oxford).

Hayward, P., Nelson-Smith, T. & Shields, C. Collins Pocket Guide. Seashore of Britain and Northern Europe. (Collins: various editions. Earlier edition (ed. Barrett & Yonge) are also useful).

Hayward, P.J. & Ryland, J.S. (1995). Handbook of the Marine Fauna of North-West Europe. (Oxford UniversityPress).

Hayward, P.J. (1988). Animals on Seaweed. Naturalists’ Handbook 9. (Richmond).

Hayward, P.J. (1994). Animals of Sandy Shores. Naturalists’ Handbook 21. (Richmond).

Hiscock, S. (1979). A field key to the British brown seaweeds. AIDGAP Field Studies Council.

Hiscock, S. (1986). A field key to the British red seaweeds. AIDGAP Field Studies Council.

Morrell, S. A key to common seaweeds. (laminated, folding guide) AIDGAP Field Studies Council.

Newell, G.E. and Newell, R.C. (1977). Marine Plankton: a practical guide. (Harper Collins).

Rainbow, P.S. (1984). An introduction to the biology of British littoral barnacles. AIDGAP Field Studies Council.

Ryland, J.S. (1986). A key for the identification of British intertidal Bryozoa. AIDGAP Field Studies Council.

Oldham, J. Guide to saltmarsh plants. (laminated, folding guide) AIDGAP Field Studies Council.

Particular groups are dealt with in the excellent Linnaean Society Monographs synopses of the British fauna, which are well illustrated and contain only the essential information for identification.

marine ecology - Ballantine, W.J. (1961). A biologically-defined exposure scale for the comparative description of rocky shores. Field Studies. Offprint no. 17: Vol. 1 (3) 1-19. Field Studies Council.

Barnes, R.S.K. and Mann, K.H. (1991). Fundamentals of Aquatic Ecology. (Blackwell).

Boyden, C.R., Crothers, J.H., Little, C. & Mettam, C. (1977). The intertidal invertebrate fauna

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39

of the Severn Estuary. Field Studies. Offprint no. 114:Vol. 4, 477-554. Field Studies Council.

Brafield, E. (1978). Life in Sandy Shores. Studies in Biology no 89. (Edward Arnold).

Crothers, J.H. (1966). Dale Fort Marine Fauna. Field Studies. Supplement to Vol. 2, 169 pages. Field Studies Council.

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