Edexcel A2 Biology Practicals (Complete)

Technician 1 of 1

Practical 5.1 Looking for patterns and correlations

Purpose

Safety

• To carry out a study on the ecology of a habitat.

Teachers/lecturers must follow their LEA/school/college policy and local rules for off-site visits, especially with regard to identification of hazards and risk assessments. Additional information is available in the DCSF guidance Health and Safety of Pupils on Educational Visits on the Teachernet website. The hazards will vary depending on the site chosen. Risk assessments will identify the risk.

The standard equipment required for carrying out an ecological study is detailed below. Additional items required to measure abiotic factors will depend on the site selected and type of study being undertaken. Requirements per student or group of students

Notes

A 50 m tape measure. If this is not available a 10–20 m tape measure can be used

A piece of rope with 0.5 m intervals marked on it will also work. Two will be needed if a grid layout is to be used.

A quadrat

Either square or point quadrats can be used. Square frame quadrats with subdivisions are useful. The ‘standard’ square frame quadrat is 50 cm 3 50 cm, but smaller and larger ones can be used.

Small, numbered pegs

At least 50.

A clipboard A clear plastic bag large enough to get clipboard and hand with pencil in

To protect students’ notes if it rains.

Ranging poles

The geography department may have these already.

A clinometer

The geography department may have one already.

Small plastic vials or dishes

For holding invertebrates while identifying them on site.

A key to organisms likely to be found in the area

The Field Studies Council produces a range of excellent guides.

Access to a compass

To describe the aspect.

Access to an OS map of the area

To accurately pinpoint the site and for background knowledge.

Jam jars

For pitfall traps.

Pooter Sweep net Tullgreen and/or Baermann funnel Thermometer Whirling hygrometer Light meter

Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

151

Student 1 of 8

Practical 5.1 Looking for patterns

Purpose • To carry out a study on the ecology of a habitat.

Observing patterns Have you ever walked into a wood and noticed that the vegetation changes as you enter? Why do the bluebells only occur under the trees? Or have you been clambering over a rocky shore and spotted that the seaweeds grow in distinctive bands and that you only find mussels where the tide is far out? What causes these patterns in plant and animal distribution? When ecologists study habitats, they try to account for plant and animal distribution, correlating them to the abiotic and biotic factors that are affecting the habitat. Abiotic means ‘non-living’ and examples of abiotic factors include light intensity, slope, humidity, wind exposure, edaphic (soil) characteristics such as pH and soil moisture, and many more. Biotic means ‘living’ and examples of biotic factors include competition, grazing and predation. All species of plants and animals you encounter in the wild are well adapted to the set of conditions encountered in their usual habitat. If they weren’t they would either grow somewhere else or become extinct!

Studying patterns Look around your local habitats and spot any patterns in distribution and abundance of organisms. You don’t need to go far; you might notice something in your school grounds or the local park. You might have a look at the distribution of plants in trampled areas of the sports field or grass paths; are there any patterns? Once you have identified a pattern, think about why it might have come about. Describe the pattern and use appropriate biological ideas to suggest an explanation. Now you need to plan a fieldwork investigation to test your idea. When planning any investigation you need to: • decide what data you are going to collect • select suitable apparatus and methods • ensure you are going to collect valid and reliable data • decide how you will analyse it once collected • complete a risk assessment and decide on steps to avoid or minimise the harmful effects of any hazards • conduct a trial to inform your planning. Read the following section, which briefly mentions some of the techniques that you could use. There is more detail in Student Practical support sheet Ecological sampling (page 26). You can also look at the British Ecological Society (BES) website education pages – the students 161 section contains detailed information about sampling techniques. Your teacher may also give you details of websites that can be used to help you prepare your plan. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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2 of 8 Student

Practical 5.1 (cont.)

Looking for patterns

Completing a transect study One of the easiest patterns to spot is zonation in the vegetation and animal distribution – as you go from one place to another the vegetation and animal distribution changes. A zonation can often be explained by a gradual change (a gradient) in one or more physical or abiotic factors. A transect is often used to study zonation in vegetation or non-mobile animal distribution. A transect is a line along which systematic records can be made.

Comparing two sites Frequently ecologists may notice a distinct pattern that does not show a gradual change and may be related to one or more factors at the two sites. For example, the vegetation in one area of a field may be very different from the rest of the field, or the species found upstream and downstream of an outflow pipe discharging into a river may seem to differ. A transect may not be the best method for this type of investigation; instead sampling of each area may be more appropriate.

Procedure 1 Plan how you are going to collect reliable and valid data that will test your hypothesis. You need to make the following decisions. • The most appropriate sampling method to use (e.g. random or systematic sampling). • The position and length of any transect to use (Figure 1). You need to make sure your transect extends far enough to sample all the possible zones. • The size and number of quadrats to use, and their positioning. • The species of plants and animals you are to record – you should focus on those which will enable you to test the hypothesis under investigation. (You may need to find out more about the species concerned using secondary sources). • The method to use for measuring abundance. • The abiotic factor(s) you are going to record. Although you may be investigating the correlation between, for example, soil moisture and the distribution of plant species, there may be other factors that could affect the distribution of organisms. It is not possible to control these variables but you can measure them and take them into account when analysing your results. • The appropriate method for measuring the abiotic factor(s). • How the data will be analysed. • How to avoid or minimise any risks when completing the fieldwork. A pilot study in advance of the main data collection will help you make these decisions. peg marked ‘20’ 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30

origin peg marked ‘0’

0.5 m � 0.5 m quadrat no. 6

transect continues

tape case

Figure 1 One way of laying out a tape measure for a transect study. Quadrats are laid down at regular intervals along the tape and the abundance of species within each quadrat is recorded. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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Student 3 of 8



2 Collect the data. 3 When you have collected your data, you must present it in an appropriate way to help you identify any patterns in the data. For transect data you can draw kite diagrams by hand or use a computer programme such as FieldWorks which may be available from your teacher. 4 Analyse your data to reveal any patterns or significant differences, and explain the main relationships between species and abiotic factors using scientific knowledge. Determine if your original hypothesis was correct. If you have suitable data, you can calculate correlation coefficients between your biotic and abiotic data. For example, you can see if there is a significant positive or negative correlation between the factor you think is responsible for the pattern and the distribution of the organisms you have recorded. Remember that correlations do not prove cause and effect. 5 In your write-up interpret your results using biological principles and concepts. Support any conclusions you make with results. Discuss the limitations of your results and conclusions based upon them, and suggest modifications that you could make to the procedure.


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4 of 8 Student

Practical support sheet Ecological sampling

Why sample in ecology? In an ideal world when investigating, say, the number of dandelions in two meadows, you would count every single dandelion in each. The problem is that this might take forever and become very, very boring. So, instead, you need to take a sample. You might estimate the number of dandelions in each meadow by counting the number in several small areas and then multiplying up to calculate a value for each meadow. The idea is to maximise the usefulness of your data while minimising the effort required to collect them.

Random sampling Frequently, ecologists notice a distinct pattern that may be related to one or more factors at two sites. For example, the vegetation in one field may be very different to that in another field, or the species found under oak trees may be different to those under ash trees, or the species upstream and downstream of an outflow pipe discharging into a river may seem to differ. To make valid comparisons, samples need to be taken from both sites. If the investigator chooses where to sample, the sample will be subjective. Random sampling allows an unbiased sample to be taken.

Using a grid

3

4

position of 0.25 m2 quadrat using random numbers 2, 4

1

2

Tape measure 2

5

In a habitat, such as a meadow or heathland, tape measures put on the ground at right-angles to each other can be used to mark out a sampling area (Figure 1). Using a pair of random numbers you can locate a position within the sampling area to collect your data. The random numbers can be pulled from a set of numbers in a hat, come from random number tables, or be generated by a calculator or computer. The two numbers are used as coordinates to locate a sampling position within the area. The first random number gives the position on the first tape and the second random number gives the position on the second tape.

1

2

3

4

5

Tape measure 1 Figure 1 Using measuring tapes to define a sample area.


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Student 5 of 8

Practical support sheet (cont.)

Ecological sampling

If you are sampling fixed objects within an area, for example the area of Pleurococcus (an alga) on the shaded side of trees in a wood or the number of woodlice under rocks, you could number all the trees or rocks and then use random numbers to select which trees or rocks to sample. This sampling idea is also used when measuring the number of cells in a culture. The culture is mixed to give a reasonably uniform distribution of cells and then a known volume is placed on a haemocytometer (a special cavity slide with a ruled grid in the centre). You then count the number of cells that occur in, say, 25 squares of the grid. Because you know the dimensions of the grid squares and the depth of the liquid above the square, you can work out the volume of culture in each square, and then calculate a mean number of cells per cm3 of the culture.

Systematic sampling Random sampling may not always be appropriate. If conditions change across a habitat, for example across a rocky shore or in a sloping meadow that becomes more boggy towards one side, then systematic sampling along a transect allows the changes to be studied. A transect is effectively a line laid out across the habitat, usually using a tape measure, along which samples are taken. The sample points may be at regular intervals, say every 2 m across a field, or they may be positioned in relation to some morphological feature, such as on the ridges and in the hollows in a sand dune system.

Sampling techniques Quadrats Quadrats are used for sampling plant communities and slow moving or stationary animals, for example many of those found on rocky shores. There are two types of quadrat: a frame quadrat and a point quadrat. A frame quadrat is usually square; the most commonly used is 50 cm by 50 cm (0.25 m2) and may be subdivided into 25 smaller squares, each 10 cm by 10 cm. The abundance of organisms within the quadrat is estimated (see the section Methods of measuring abundance and Figure 3). Quadrats may be placed across the site to be sampled using random or systematic sampling methods. Throwing quadrats is not random and can be dangerous. It is important to sample enough quadrats to be representative of the site, but why do 1000 quadrats if 10 will give almost as accurate a result? To find out the optimum number of quadrats required, record the number of species in each quadrat and plot the cumulative results against number of quadrats until sampling additional quadrats does not substantially increase the number of species recorded. A point quadrat frame (Figure 2) enables pins to be lowered onto the vegetation below. Each species touched is recorded as a hit. The percentage cover for a particular species is calculated using the equation: hits     3 100 % cover 5 ____________   hits 1 misses Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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6 of 8 Student


Ecological sampling

knitting needle

holes metal spike (such as a tent peg) inserted in ground multiple hit

Figure 2 A point quadrat frame. Each plant species touched by the needle is recorded.

Methods of measuring abundance Density Count the number of individuals in several quadrats and take the mean to give number per unit area, for example per metre squared (m22). In many plant species (e.g. grasses) it is very difficult to distinguish individual plants, so measuring density is not possible.

Frequency Frequency is the number or percentage of sampling units in which a particular species occurs. This avoids having to count the number of individuals. If clover was recorded in 10 of the 25 squares that make up a 0.25 m2 quadrat frame, the percentage frequency would be 40%. You need to be consistent when determining presence or absence in a sampling unit. For example, you might decide that only plants rooted in the square are counted, or you might decide that any plant or animal in the quadrat is counted including any that touch or overhang the quadrat.

Percentage cover This is the percentage of the ground covered by a species within the sampling unit. Count the number of squares within the quadrat that the plant completely covers, then count those that are only partly covered and estimate the total number of full squares that would be completely covered by that species.

Estimating animal populations Quadrats cannot be used for mobile animals as these don’t stay in the quadrats. A variety of different nets and traps need to be used. Animals that occur on the soil surface may be sampled using a pitfall trap (Figure 3). Those in vegetation can be sampled using a pooter directly or indirectly (after being knocked from the vegetation onto a white sheet). Insects and other small invertebrates found in leaf litter can be collected using a Tullgren funnel. Mark–release methods can also be used.


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Student 7 of 8


Ecological sampling

• Pitfall trap for sampling arthropods flat stone prevents rainfall filling trap stick support

• Tullgren funnel for collecting organisms from the soil or leaf litter

25 W bulb soil sample 16 mesh flour sieve

jam jar sunk into soil

ground slopes away from trap for drainage

bait of meat or ripe fruit polythene funnel • Pooter for collecting insects glass collecting tube

clear plastic tube

air and arthropods drawn into tube cork or rubber bung specimen tube where arthropods collect

Organisms move away from the heat and light, falling into the jar.

glass mouthpiece

80% alcohol

gauze covering tube opening air sucked through mouthpiece

• Alternatively a muslin bag of soil surrounded by water can be used to collect living organisms. This is a Baermann funnel.

60 W bulb • Sweep net This net is swept through low-growing vegetation, collecting any animals in the mesh net.

rod for supporting bag soil sample in muslin bag water glass funnel rubber tubing

clip

beaker

Figure 3 Net and traps for sampling animals.


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8 of 8 Student


Ecological sampling

Measuring abiotic factors when sampling the environment Angle of slope Use a clinometer.

Aspect Use a compass.

Temperature Use a thermometer or temperature probe, but be aware that the time of day can influence the values obtained, as will cloud cover. The thermometer or probe should be placed in the same position each time a measurement is made to allow valid comparison of measurements.

Light Use a light meter. Light readings can vary widely with time of day and cloud cover. It is better to take all measurements over a short period or take regular readings over extended periods using a datalogger.

Oxygen concentration In aquatic systems, oxygen probes can be used to measure oxygen concentration.

Humidity Relative humidity can be measured using a whirling hygrometer. It needs to be spun for 60 seconds just above the vegetation before readings are taken from the wet and dry thermometer and used to determine the humidity from a calibration scale.

Conductivity The ability of a water sample to carry an electric current gives a measure of the dissolved mineral salts. The conductivity of pure water is zero; increasing ion concentration raises the conductivity.

Soil water A sample of soil is dried at 110 ºC until there is no further loss in mass. The % soil moisture can be calculated using the equation: mass of fresh soil 2 mass of dry soil  3 100 % soil moisture 5 ________________________________            mass of fresh soil

Soil organic matter A dry soil sample of known mass is heated in a crucible for 15 minutes to burn off all the organic matter. The mass is re-measured after the soil sample has cooled. The % soil organic matter is calculated using the equation: mass of dry soil 2 mass of burnt soil  3 100 % organic matter in soil 5  ________________________________          mass of dry soil

pH Universal Indicator or a pH meter can be used to test pH after mixing a soil sample with water. If using Universal indicator in the field, it is best to use a proper soil testing kit that contains some long glass tubes, with lines engraved on the sides, to show levels for adding soil and chemicals. First, 1 cm3 of soil is shaken with distilled water before adding one spatula of barium sulphate (low hazard). This helps to flocculate (settle) the clay fraction, which is important as clay particles are very small and will otherwise cloud the water for days. Then 1 cm3 of pH indicator solution is added and the pH recorded after the contents of the tubes have been allowed to settle. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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Teacher/Lecturer 1 of 3

Practical 5.1 Looking for patterns

Purpose

Safety

• To carry out a study on the ecology of a habitat.

Teachers/lecturers must follow their LEA/school/college policy and local rules for off-site visits, especially with regard to identification of hazards and risk assessments. Additional information is available in the DCSF guidance Health and Safety of Pupils on Educational Visits on the Teachernet website. The hazards will vary depending on the site chosen. Risk assessments will identify the risk.

Notes on the procedure Students need to carry out a study on the ecology of a habitat to produce valid and reliable data (including the use of quadrats and transects to assess the abundance and distribution of organisms and the measurement of abiotic factors). This activity outlines how to approach a fieldwork investigation. It briefly mentions some techniques that might be used. It does not attempt to provide detailed accounts of the techniques. Additional details are provided on the Student Practical support sheet Ecological sampling (page 26) and there is a wealth of information on the British Ecological Society student 161 education web pages about sampling techniques. There are also examples of projects with data and virtual tours of the rocky shore and sand dune habitats. The Field Studies Council website also has online resources for students including information on different urban ecosystems. The methods used will depend on the habitat and factors under investigation. Any suitable methods could be used that give students the opportunity to have first-hand experience of field data collection. The school/college grounds can be a valuable resource. Other activities could be covered in a fieldwork context; for example, feeding relationships and the transfer of energy through ecosystems, or succession, could be investigated through a specific habitat. Fieldwork can be used as students’ investigation for A2 coursework. Teachers/ lecturers and students should ensure that the proposed investigation meets the assessment criteria. When planning coursework for Unit 6 there is a full description of the requirements illustrated with exemplars in the Tutor support material for Unit 6 on the Edexcel website.

Select a site carefully The site selected should show some clear relationships. Some examples where transects can be used: • a playing field or grassy area from an area of high trampling to an area of low trampling. The Field Studies Council online resources include a case study on the distribution of ribwort plantain and greater plantain in trampled and non-trampled areas.


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2 of 3 Teacher/Lecturer



• a woodland margin – passing from a field or other example of grazed or mown grassland, through brambles into a wood. The key gradient is likely to be light intensity – but it may not be the only one. • a sand dune system from the shore across dunes into grassland and scrub as you go further inland. In this case the key factors could be soil moisture, soil stability and organic matter, although succession is also involved here. • a rocky shore from the low tide mark to the top of the beach. The key factor is the proportion of time a part of the shore is left exposed to the air and to desiccation when the tide is out. • a geological boundary between two types of rock, such as between limestone and millstone grit. Some examples where two sites can be compared: • grazed and ungrazed grassland • mowed and unmowed grassland • trampled and untrampled grassland • fertilised and unfertilised lawns • shaded and sunny sites • fast- and slow-flowing streams • chalk and sandy soil sites • understories of beech and oak woodlands. The two sites could be compared by random sampling.

Identification – don’t panic! Identification is not as big a problem as you may think. The only species students need focus on are those which would support the hypothesis under investigation. Trying to identify each species in a quadrat down to the tiniest piece of moss can be a poor use of time and can actually prevent students from ‘seeing the wood for the trees’. A transect at a woodland margin could be successfully done recording only tree cover, grass, brambles, nettles, ivy, bare ground, wild garlic (with give-away smell) and bluebells, together with good light meter readings and soil pH (often showing no pattern), especially if you start to consider something like leaf surface area of brambles at the same time. But the principles are: • select a site where students can cope with the species identification • before teaching students to identify another species, ask yourself ‘What is the ecological point of teaching this?’ • make use of expert help if available – this could include a Field Study Centre. The Field Studies Council produces excellent laminated cards to aid with field identification in particular habitats. The students will be relying on you for identification so careful preparation is important. You may like to produce a record sheet that includes the species you want to focus on. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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Class organisation Students can work in groups if they are not using this investigation for their A2 coursework. The smaller the group the more ‘ownership’ by individuals, whilst the larger the group the more quadrats can be recorded and the bigger and statistically more meaningful the picture that can be produced. A good group size is six – working as three pairs. It helps if the whole group can have access to a computer as soon as possible after collecting the data and to a full set of equipment during data collection. Waiting to borrow equipment from other groups wastes a lot of time and loses momentum. Pace is also important – it is possible to be too slow and nit-picking about accuracy, in which case the students become bored and lose sight of the bigger picture. It is also possible for the students to feel it is somehow ‘efficient’ to get the job done as quickly as possible, subsequently data are produced that are so inaccurate and incomplete that they do not produce reliable results.

FieldWorks: an invaluable IT resource FieldWorks is available on CD and produced by Interpretive Solutions, Hallsannery Field Centre, Bideford, Devon EX39 5HE. It benefits from having grown out of the learning experiences of many students. Students can enter their own data, and present these using kite diagrams or other formats. It also contains information about species and habitats including rocky shores, sand dune, salt marsh, moorland, woodlands and fresh water. The data can be subjected to statistical analysis using correlation coefficients (Pearson’s parametric or Spearman rank non-parametric), particularly useful with transects. The programme can also calculate species diversity indices and carry out t-tests and z-tests. Other statistical packages are available for use in ecology from software companies and other suppliers.

Useful references Jones C. (1998) Fieldwork sampling – animals. Biological Sciences Review, 10(4), 23–25. Jones C. (1998) Fieldwork sampling – plants. Biological Sciences Review, 10(5), 6–8. Williams G. (1987) Techniques and fieldwork in ecology. London: HarperCollins. Field Studies Council identification sheets and booklets are very good. These can be obtained from: FSC Publications, Preston Montford, Montford Bridge, Shrewsbury SY4 1HW.


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1 of 1 Technician

Practical 5.2 The effect of temperature on the hatching success of brine shrimps Purpose • To investigate the effect of temperature on the hatching success of brine shrimps. • To develop certain experimental skills, namely considering the ethical issues arising from the use of living organisms, presenting results, producing reliable results, identifying trends in data and drawing valid conclusions.

For detailed guidance on the care and breeding of brine shrimps see the publication Brine Shrimp Ecology by Michael Dockery and Stephen Tomkins, available from: Homerton Brine Shrimp Project, Dept of Biological Sciences, Homerton College, Cambridge CB2 2PH Tel: 01223 507175. Price from Blades Biological, £48.26 (2009) including post and packing, brine shrimp eggs and innoculum. For more details see the teacher area of the British Ecological Society website.

General note This practical takes place over several days. The students set up the beakers with eggs to hatch on the first day. On subsequent days they count the number of eggs that have hatched. Requirements per student or group of students

Notes

Brine shrimp egg cysts

Available from pet shops. There are approximately 24 000 egg cysts per gram so only tiny quantities are required. Brine shrimps will breed and produce cysts that can be collected. They adhere to the sides of an aquarium tank.

100 cm3 beakers (one for each temperature to be tested) 100 cm3 de-chlorinated water for each treatment

Tap water can be de-chlorinated by leaving it to stand for 48 hours.

3

40 cm beaker of salt water 2 g sea salt for each treatment

Students could be supplied with the salt water already prepared.

Stirring rod Access to refrigerator Access to water baths or incubators (one for each temperature to be investigated)

A range of temperatures between 5 and 35 °C is recommended. The optimum for hatching is 28 °C.

White A4 sheet of paper Sheet of graph paper 3 cm × 4 cm Magnifying glass Pair of forceps Bright light

A lamp or light from one side. For counting larvae on the second day.

Fine glass pipette

For counting larvae on the second day.

Small beaker of salt water

For counting larvae on the second day.

Large beaker or tank of salt water 1 substrate and algae, set up well in advance. Details in the Dockery and Tomkins book

To hold the brine shrimps after hatching. This tank can be maintained for subsequent investigations of brine shrimps.


152

Student 1 of 2

Practical 5.2 The effect of temperature on the hatching success of brine shrimps Purpose • To investigate the effect of temperature on the hatching success of brine shrimps. • To develop certain experimental skills, namely considering the ethical issues arising from the use of living organisms, presenting results, producing reliable results, identifying trends in data and drawing valid conclusions.

Brine shrimps Brine shrimps are small, saltwater crustaceans; the adults are about 8 mm in length. They are relatively easy to keep in the laboratory and will produce dormant egg cysts that hatch to produce young shrimp larvae. first antenna male (blue/green)

female (brownish red) 1 mm eggs

second antenna Drawings to show features of brine shrimps

Procedure You will need: • Brine shrimp egg cysts • 2 g sea salt for each treatment • 100 cm3 de-chlorinated water for each treatment • 40 cm3 beaker of salt water • 100 cm3 beakers (one for each temperature to be tested) • Water baths or incubators (one for each temperature to be investigated)

• • • • • • • •

Stirring rod Magnifying glass Pair of forceps Fine glass pipette Bright light Access to refrigerator Sheet of A4 white paper Sheet of graph paper 3 cm 3 4 cm

1 Decide on a range of temperatures from 5 °C to 35 °C to be tested. 2 Place 2 g of sea salt into a 100 cm3 beaker. 3 Add 100 cm3 of de-chlorinated water and stir until the salt completely dissolves. 4 Label the beaker with sea salt and the temperature at which it will be incubated. 5 Place a tiny pinch of egg cysts onto a large sheet of white paper. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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2 of 2 Student


The effect of temperature on the hatching success

of brine shrimps 6 Wet the piece of graph paper using a few drops of salt water. Dab the paper onto the white sheet to pick up approximately 40 eggs. This will look like a tiny shake of pepper. Use a magnifying glass to count the eggs. Cut the graph paper so that there are exactly 40 eggs. 7 Put the paper with the 40 eggs into the beaker (eggs-side down). After 3 minutes, use a pair of forceps to gently remove the paper, making sure that all the egg cysts have washed off into the water. 8 Repeat steps 2 to 7 for all the temperatures that are to be investigated. 9 If possible replicate the treatments. 10 Incubate the beakers at the appropriate temperatures, controlling exposure to light as far as possible. 11 The next day count the number of hatched larvae in each of the beakers. To do this, place a bright light next to the beaker. Any larvae will swim towards the light. Using a fine glass pipette catch the brine shrimps and place them in a small beaker of salt water. (It may be easier if the pipette is reversed with the tip inserted into the teat, providing a wider bore to take up the shrimp.) Repeat the counting daily for several days. Brine shrimps are very delicate and care must be taken when handling them. Finally, discuss with your teacher the best method for disposing of the brine shrimps. 12 Record the number of larvae that have successfully hatched at each temperature. 13 Write up your experiment making sure your report includes: • a discussion of any health and safety precautions taken • comments on the ethical issues arising from the use of living organisms • results presented in the most appropriate way • an explanation of any patterns in the data using evidence from the data and your own biological knowledge • comments on how valid your conclusion is • comments on how you ensured that the results obtained in this experiment were valid and reliable • suggestions for how you could have made your results more reliable. To find out more about brine shrimps, visit the British Ecological Society website.


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Practical 5.2 The effect of temperature on the hatching success of brine shrimps Purpose • To investigate the effect of temperature on the hatching success of brine shrimp. • To develop certain experimental skills, namely considering the ethical issues arising from the use of living organisms, presenting results, producing reliable results, identifying trends in data and drawing valid conclusions.

Notes on the procedure If students are given basic information on maintaining brine shrimps this activity could be planned before they are given the Student sheet. The need to keep conditions other than temperature constant should be appreciated. Brine shrimps hatch in salt water that is 2 to 5% salt (optimum 2.8%) and pH 8.5. Oxygen must be present. Light is an added (but not essential) factor for hatching. Datalogging could be used to check that the conditions within each of the treatments are maintained at a constant level. This experiment could be completed by small groups of students. If each group completed the range of temperatures then the error between replicates could be investigated. Brine shrimps will hatch in 24 hours at temperatures between 25 and 30 °C with an optimum of 28 °C. This activity can be used to highlight experimental and investigative assessment objectives, in particular the ethical issues arising from the use of living organisms (including the disposal of the brine shrimps after the activity) and for the environment, and the need to identify both dependent and independent variables and, where possible, control or allow for them. The method suggested provides precise measurements. The need for valid, reliable results should be considered and the random nature of error could be investigated. The practical procedure and technical notes are based on an investigation that appears in the British Ecological Society publication Brine Shrimp Ecology by Michael Dockery and Stephen Tomkins. This book contains detailed information on the care and breeding of brine shrimps. It is available from the British Ecological Society or from: Homerton Brine Shrimp Project Dept of Biological Sciences Homerton College Cambridge CB2 2PH Tel: 01223 507175 Price from Blades Biological, £48.26 (2009) including post and packing, brine shrimp eggs and innoculum. For more details, including the preparation of a salt water aquarium in which to keep the brine shrimps, see the teacher area of the British Ecological Society website.


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Student 1 of 1

Practical 6.1 DNA gel electrophoresis

Purpose

Safety

• To use gel electrophoresis to separate DNA fragments of different sizes.

If you undertake gel electrophoresis make sure you are aware of the hazards and follow the instructions of your teacher very carefully.

Procedure You may have the opportunity to complete experimental work using restriction enzymes and gel electrophoresis or you may use the simulation of this.


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Practical 6.1 DNA gel electrophoresis

Purpose

Safety

• To use gel electrophoresis to separate DNA fragments of different sizes.

If gel electrophoresis is to be used a full risk assessment should be obtained for the procedure and carefully followed by both staff and students.

Restriction enzymes The use of restriction enzymes to cut DNA and electrophoresis to separate the resulting fragments is possible using equipment available from NCBE (National Centre for Biotechnology Education) and Bio-Rad. Protocols for these practicals as pdf files can be downloaded from their websites. NCBE produce an electrophoresis base unit and a lambda DNA kit. Together these contain all necessary gel electrophoresis apparatus, dried lambda DNA, three dried restriction enzymes and both student and technical guides. The kit includes microsyringes and gel tanks but not batteries needed as a power supply. The kit for eight electrophoresis stations (16 runs) cost £52 in 2009. The lambda protocol module, for use with eight students, cost £90 in 2009. Have a look at the technical guides by downloading the pdf files from the NCBE website. The NCBE publication Illuminating DNA also contains protocols for experiments using restriction enzymes and electrophoresis. This publication can also be viewed on the NCBE website. NCBE has developed a genetic screening simulation, Nature’s dice. In this practical simulation, the inheritance of a single gene that is involved in a particular genetic trait is investigated. The kit contains 24 DNA samples donated by a large family who are affected by the genetic condition. Each student is given one or more of these DNA samples and they have to detect which allele is present. They cut the DNA with a restriction enzyme and examine the resulting DNA fragments by electrophoresis. The genetic data obtained is then combined with the family tree to look at how the genetic trait is inherited. This activity would provide an excellent alternative to the standard gel electrophoresis practical suggested above. Further information about the kit and cost of materials is on the website listed in the weblinks section for this activity. Bio-Rad produces a Restriction Digestion and analysis of lambda DNA kit. This contains dried lambda DNA, three restriction enzymes, buffers, microtubes and both Student and Teacher’s guides. It does not include the micropipette, electrophoresis cell or power supply. The kit for eight stations (eight runs) cost £86 in 2009; the cell and power supply needed to run the gels cost an additional £4001 but can be borrowed from some ITT centres (see below). You can have a look at the protocol by downloading the pdf file from the Bio-Rad website. Their website contains more detail and alternative kits. Go to the site and then click the Life Science education icon. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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DNA gel electrophoresis

Pfizer and Bio-Rad have donated biotechnology equipment to 31 ITT centres around the country as part of a National Year of Science project. Each centre holds equipment that can be used to conduct biotechnology practicals for 25 students at a time. They have all the equipment necessary for completing DNA fingerprinting, bacterial transformation (genetically altering bacteria with a bioluminescent jellyfish gene), purification of a useful protein, and the polymerase chain reaction (PCR). It is envisaged that this equipment will be used for hands-on practical training of student science teachers. There are also plans for loan schemes to be established to enable local schools to borrow the equipment. To find out more about this project contact the BioEducation Project Manager at Bio-Rad Laboratories (details can be found on the BioRad website). Both organisations produce replacement equipment for their kits and also supply the components separately. Note that some of the Bio-Rad manuals accompanying their kits (unless recently revised) may not carry full and appropriate health and safety guidance applicable in the UK (they were written for a US audience). Where the instructions conflict with good practice that is standard in the UK, additional precautions should be taken. It is most likely that gel electrophoresis equipment using batteries or a low-voltage supply is used. This is safe if the voltage is less than 40 V DC. If equipment is used that is operated by a power supply at a greater voltage, it is essential that the equipment is designed so that it is impossible to make skin contact with the electrodes or electrolyte when an electric current is flowing. This is usually achieved by a suitable design of lid for the gel tank which only permits a current to flow when the lid is in place. Also, although it is very unlikely to be needed, ethidium bromide, which is very toxic, is not normally recommended for use in schools. For general CLEAPSS guidance on electrophoresis, schools should consult section 11.1.7 of the Laboratory Handbook.


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1 of 1 Student

Practical 6.2 DNA amplification using PCR

Purpose

Safety

• To use PCR to amplify DNA.

If you undertake PCR make sure you are aware of the hazards and follow the instructions of your teacher very carefully.

Practical PCR Scientists in forensics laboratories carry out the polymerase chain reaction (PCR) using a machine called an automated thermal cycler. This is a programmable heating unit in which the DNA to be amplified is incubated in a buffer solution with thermo-stable DNA polymerase, primers and deoxyribonucleotides. The unit maintains the cyclical sequence of temperatures for the PCR process. Your school or college may be lucky enough to possess a thermal cycler but it is possible to carry out PCR without them, using three separate thermostatically controlled water baths. You simply have to move the DNA sample from bath to bath – and complete 30 cycles! You need a stopwatch, good teamwork and some sort of protection from the steam coming off the hottest bath. Having amplified the short tandem repeat sequences within your DNA sample, you will then separate out the fragments using gel electrophoresis (see Practical 6.1). Comparing the position of the bands on the gels to a standard or reference you will be able to draw conclusions about the DNA sample you started with. You will follow a practical protocol supplied by the company that produces the equipment and reagents your school or college has purchased.


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Practical 6.2 DNA amplification using PCR

Purpose

Safety

• To use PCR to amplify DNA.

If PCR is to be used a full risk assessment should be obtained for the procedure and carefully followed by both staff and students.

PCR It is possible to carry out practical PCR using equipment available from a number of suppliers including NCBE (National Centre for Biotechnology Education), Bio-Rad and Edvotek Europe. Although all organisations supply automated PCR thermal cyclers, they are not absolutely necessary. Instead, PCR can be undertaken using three separate thermostatically controlled water baths, although care must be taken with the hottest as a risk of scalding exists. Note that during Science Year (September 2001–July 2002) many schools, colleges and initial teacher training institutions received free PCR equipment. Student and teacher/technician protocols for the PCR practicals can be downloaded as pdf files from the suppliers’ websites. Equipment can be purchased either in class sets or individually, with consumables also available in class-sized batches. All three organisations listed offer training courses, frequently in association with other institutions like botanic gardens or science centres, giving teachers and technicians the opportunity to carry out the practicals themselves. There are social and ethical considerations to take into account when using DNA derived from students for PCR. Although the suppliers should have chosen STR sequences that have no biological significance, it may be socially and ethically less challenging to use plant DNA as the sample. But working with one’s own DNA can be uniquely motivating!


13



DNA amplification using PCR

Contact details and examples of the protocols offered by NCBE, Bio-Rad and Edvotek Europe: Organisation

Example of suitable protocols

NCBE National Centre for Biotechnology Education Science and Technology Centre The University of Reading Whiteknights Reading, RG6 6BZ www.ncbe.reading.ac.uk 0118 987 3743 [email protected]

Amplifying lambda DNA (from the Illuminating DNA booklet) Investigating plant evolution Analysing mitochondrial DNA. This practical requires a microcentrifuge and the protocol is not downloadable from the NCBE website. It is based on a protocol developed for a DNA workshop at the Cold Spring Harbor Laboratory. It can be obtained direct from NCBE.

Bio-Rad Bio-Rad Laboratories Ltd. Bio-Rad House Maxted Road Hemel Hempstead Hertfordshire HP2 7DX www.biorad.com/ Follow the links to Life Science education, then About Biotechnology Explorer Freephone: 0800 181134

Crime scene investigator – PCR basics PV92 PCR infomatics kit GMO investigator kit (only useful if GM foodstuffs are easily available)

Edvotek Europe The Biotechnology Education Company PO Box 280 Hertford SG13 9DG www.edvotek.co.uk Follow links to Experiments and then Polymerase chain reaction [email protected]

332 Mitochondrial DNA analysis using PCR


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Technician 1 of 1

Practical 6.3 Which antibiotic is most effective?

Purpose • To investigate the effect of different antibiotics on bacteria.

This experiment could be done with antiseptics rather than antibiotics. Paper discs produced with a hole punch are autoclaved, then dipped in a range of antiseptics. CLEAPSS guidance on microbiology is in section 15.2 of the Laboratory Handbook. Sterilisation guidance is in section 15.12. Requirements per student or group of students

Notes

Agar plates seeded with bacteria or equipment for their preparation (i.e. access to one of two broth cultures of known bacteria; Universal McCartney bottle of nutrient agar; sterile Petri dish; sterile pipette)

Suitable bacteria are listed in various catalogues. Phillip Harris use E. coli (K12 strain) and Staphylococcus albus in their kit. The Student sheet with instructions for preparation of seeded agar plates, ‘Pouring agar plates’, can be found in Edexcel AS Biology Practical 4.3. Safety The bacterial culture could present a biohazard and should be handled and disposed of in accordance with current guidelines on microbiology in schools. Disposal of agar plates and other used equipment, including forceps, must be in accordance with current best practice. Autoclaving is the preferred method of disposal over any chemical disinfection method.

Bunsen burner Bench spray of disinfectant

1% Virkon (preferred) or equivalent.

Soap or handwash

This does not need to be bactericidal.

Paper towels Marker pen for marking Petri dishes Autoclaved after placing in cotton wool stoppered boiling tube.

Forceps (metal) A Mast ring or separate antibiotic discs

Separate antibiotic discs are usually cheaper, but it is impossible to obtain more than two types of antibiotics using these.

Adhesive tape Space to keep dishes at room temperature

This will be needed for at least 48 hours.

Eye protection

Not essential.


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Student 1 of 2


Purpose

Safety

• To investigate the effect of different antibiotics on bacteria.

Wear eye protection. The microorganisms are a potential biological hazard. Use aseptic techniques when transferring the bacteria to the Petri dishes. Clean the bench with antibacterial disinfectant. Do NOT open the Petri dishes once they have been incubated.

Introduction When a bacterial infection is diagnosed it is useful to be able to tell to which antibiotics it is most susceptible. In some cases this information is known, but in other cases tests need to be carried out to find out which antibiotic will be most effective. In this activity you will be testing the effectiveness of several types of antibiotics on bacteria. The standard method of doing this is to put discs of chromatography blotting paper soaked in the various antibiotics onto an agar plate that has been inoculated with the bacteria. Alternatively a Mast ring (a ring of paper with several ‘arms’, each treated with a different antibiotic) can be used.

Procedure You will need: • Agar plate seeded with a known bacterium • Bunsen burner • Bench spray of disinfectant, 1% Virkon or equivalent • Soap or handwash • Paper towels

• Marker pen • Autoclaved forceps • Mast ring or antibiotic-impregnated paper discs • Adhesive tape • Eye protection

1 Wash your hands with the soap or handwash. Spray the working area thoroughly with the disinfectant spray. Leave for at least 10 minutes, then wipe with a paper towel. 2 Work very close to a lit Bunsen burner. Prepare an agar plate seeded with bacteria. This may have already been done for you. If not, follow the instructions in the section ‘Pouring agar plates’ in Practical 4.3 Edexcel AS Biology. Label the Petri dish on the base at the edge with your name, the date and the type of bacterium it is inoculated with. 3 Flame the forceps and then use them to pick up an antibiotic disc or Mast ring. Raise the lid of the Petri dish and place the Mast ring firmly in the centre of the agar; if individual discs are used they will need to be spaced evenly around the dish. 4 Tape the dish securely with two pieces of adhesive tape (but do not seal it completely), then keep it upside down at room temperature for 48 hours. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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Which antibiotic is most effective?

5 Wash your hands with soap or handwash and clean the bench again using the Virkon spray. 6 After incubation, look carefully at the plate but do not open it. Where bacteria have grown the plate will look opaque, but where the antibiotics have inhibited growth, clear zones called inhibition zones will be seen. Measure the diameter of the inhibition zones in millimetres and use this information to decide which antibiotic is most effective at inhibiting the growth of the bacterium. 7 Collect data from other members of the class who used the other bacterial cultures. 8 Write a brief report of the results, comparing the different antibiotics and the effects on the different bacterial cultures.

Questions 1 Are the inhibition zones circular? If not, what is a sensible measuring strategy? 2 What factors determine the diameter of the inhibition zones? 3 If class data are shared: a what is the overall spread of the data b do all individual results show the same trends – if not, why not, and how could this variability be represented on your graphs? 4 If you were working in a hospital laboratory, and you had just carried out this test on bacteria isolated from sick patients, would you always choose the antibiotic that gave the biggest inhibition zone? Are there any other factors you would need to consider?


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Purpose

Safety

• To investigate the effect of different antibiotics on bacteria.

Eye protection is not necessarily required during microbial work, though it is good standard practice. The microorganisms are a potential biological hazard. Use aseptic techniques when transferring the bacteria to the Petri dishes. Clean the bench with antibacterial disinfectant. Do NOT open the Petri dishes once they have been incubated.

Notes on the procedure The aim of the practical is to show students a method of determining bacterial sensitivity to different antibiotics and to give them practice at using aseptic techniques. This could be done with antibiotic discs or the same technique could be used with antiseptics. The Student sheet suggests using different bacterial species but the activity could be done using just one species. However, using different species should illustrate that they are not all equally susceptible. Although allergic response by patients is considered in the questions, where antibiotics are already impregnated on to discs, the risk for students with allergic responses is not great. Discs are not handled directly and no airborne dust is created by the discs. The behaviour of students must be considered. If they might attempt to lift a piece of tape from an incubated dish, the agar plates should be sealed around the circumference after incubation but before being returned for observation. Also, to prevent agar plates being removed from the lab, count all plates back in again before students are allowed to leave.

Answers 1 Choice of strategy may vary, but it must be clear and easy to carry out and produce reliable results. 2 The rate of diffusion of the antibiotic will be influenced by the size of molecule, its concentration and the potency of the antibiotic. If the antibiotic is effective at lower concentrations the circle will be larger (all other things being equal). Gram-positive and Gram-negative bacteria respond differently to antibiotics. 3 Responses will depend on the data, but should show a clear understanding of the concepts of accuracy, validity and/or reliability when explaining variation. Suggestions for the presentation of variation in graphs may also vary but need to identify anomalous results clearly so that conclusion are obviously based on reliable results. 4 You may have to consider whether the patient is allergic to any of the antibiotics. For example, allergy to penicillin is not uncommon. You may consider the state of the patient’s immune system. In patients with a weakened immune system you would not want to use a bacteriostatic antibiotic, that is, one that stops bacterial reproduction but does not kill the bacteria. Some antibiotics can be used together and produce a larger effect when combined than if administered separately. This is known as synergism. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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1 of 1 Technician

Practical 7.1 Measuring the rate of oxygen uptake

Purpose • To demonstrate the uptake of oxygen in respiration. • To measure the rate at which an organism respires.

Requirements per student or group of students

Notes

Respirometer

See the diagram on the Student sheet. If a pipette is used, the scale shown will not be needed. Ideally the syringes are attached with a three-way tap. If these are not available a rubber tube and clip can be used. U-tube respirometers would be even better, if there is a class set available. See also section 15.10 on respirometers in the CLEAPS Laboratory Handbook for details of other (bulk) suppliers.

5 g of an actively respiring organism

Use actively germinating peas, beans or other seeds, or maggots, or woodlice.

Roughly a tablespoon of soda lime

To absorb the carbon dioxide. Safety Soda lime is corrosive, but much less of a hazard than solutions of potassium or sodium hydroxide. Even so, eye protection is needed when handling the soda lime. Do not handle directly: use a spatula. Soda lime can sometimes be ’dusty’. To avoid exposing invertebrates to corrosive dust, take the soda lime outside and pour from beaker to beaker to blow the dust away. Position yourself so that you cannot inhale the dust during this activity.

About 2 cm3 of coloured liquid

e.g. water and food colour or equivalent.

Dropping pipette

Or 1 cm3 syringe plus hypodermic needle (in cover until in use) if appropriate to students.

Permanent OHT marker, or chinagraph pencils

For marking the position of the coloured liquid.

Solvent to remove the marker Small amount of cotton wool to wipe pipette Stopclock Eye protection

Respirometers A respirometer is shown on the Student sheet. Many schools and colleges have at least one of the U-tube respirometers (Figure 7.1.4 on page 131 of the A2 textbook). These can be used but can be a lot more fiddly, and the connections often leak. If the apparatus works, the respiring organisms use up the oxygen and give off CO2. The CO2 is absorbed by the soda lime. This means there is less air in the test tube so the liquid gets sucked towards the tube with the organisms in it. If it does not work there is usually an air leak somewhere, or possibly the organisms are too cold, or dead!


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Student 1 of 2


Purpose

Safety

• To demonstrate the uptake of oxygen in respiration. • To measure the rate at which an organism respires.

Wear eye protection when handling soda lime. Soda lime is corrosive. Do not handle directly: use a spatula.

Respirometers Respirometers range from relatively simple pieces of equipment used in school science labs with seeds or invertebrates, to elaborate devices the size of a room used to measure respiration rates in humans living near-normal lives over a period of several days. In this practical you will be using a very simple respirometer, while considering the advantages of some of the slightly more complex ones.

Procedure You will need: • Respirometer (see diagram below) • 5 g of an actively respiring organism

• • • •

Soda lime Coloured liquid Dropping pipette Permanent OHT marker pen

• • • •

Solvent (to remove the marker) Cotton wool Stopclock Eye protection

1 Assemble the apparatus as shown in the diagram below.

syringe

scale

three-way tap glass tubing

1 cm3 pipette or glass tube

coloured liquid

small organisms gauze soda lime A simple respirometer

2 Place 5 g of maggots or peas into the test tube and replace the bung.


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Measuring the rate of oxygen uptake

3 Introduce a drop of marker fluid into the pipette or glass tube using a dropping pipette. Open the connection (three-way tap) to the syringe and move the fluid to a convenient place on the pipette (i.e. towards the end of the scale that is furthest from the test tube). 4 Mark the starting position of the fluid on the pipette tube with a permanent OHT pen. 5 Isolate the respirometer by closing the connection to the syringe and the atmosphere and immediately start the stopclock. Mark the position of the fluid on the pipette at 1 minute intervals for 5 minutes. 6 At the end of 5 minutes open the connection to the outside air. 7 Measure the distance travelled by the liquid during each minute (the distance from one mark to the next on your pipette).

If your tube does not have volumes marked onto it you will need to convert the distance moved into volume of oxygen used. (Remember the volume used 5 pr2 3 distance moved, where r 5 the radius of the hole in the pipette.)

9 Record your results in a suitable table. 10 Calculate the mean rate of oxygen uptake during the 5 minutes.

Questions 1 Why did the liquid move? Explain in detail what happens to the oxygen molecules, the carbon dioxide molecules and the pressure in the tube. 2 It would have been better to set up a second, control tube that did not contain living organisms but had everything else the same. a What could cause a movement of the liquid in the control tube towards the respirometer? b What could cause a movement of the liquid in the control tube away from the respirometer? c What could you do to correct your estimate of oxygen uptake if the liquid in the control had moved too?

Extension 3 The diagram below and Figure 7.24 on page 148 of A2 Biology show two other types of respirometer. What advantages and disadvantages do these have compared to the one you are using? soda lime

drop of liquid

wire mesh

organism to be studied

capillary tube

A very simple respirometer

4 Design an experiment to investigate the effect of different temperatures on the rate of oxygen uptake in maggots. Remember that the maggots will need time to acclimatise to each new temperature. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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Purpose

Safety

• To demonstrate the uptake of oxygen in respiration. • To measure the rate at which an organism respires.

Wear eye protection when handling soda lime. Soda lime is corrosive. Do not handle directly: use a spatula.

Notes on the procedure Rate of respiration is not a learning outcome in the specification. The respirometer shown in the diagram is a very simple one; more complex ones (e.g. U-tubes) can be used if available. Question 1 can be used as a summary activity for student understanding of respiration if students are asked to complete it in as much detail as possible and to state the source of carbon dioxide as well as its fate. The choice of what respiring organisms to put in the tubes is left to you. Germinating peas, maggots or woodlice are commonly used. If you are using pipettes there is no need to do any volume calculations; students just read the change in volume off the scale on the pipettes. If you are using thick-walled capillary tubing it is worth reminding students of the formula for working out the volume of oxygen used, and explaining it again. volume of oxygen used 5 pr2 3 distance moved where r 5 the radius of the capillary tubing or pipette. Three-way taps can cause confusion with some students. A diagram of how the three-way tap works is shown in the diagram below. Putting up an OHT of this diagram during the practical can help. 1

2

respirometer open to syringe

3

respirometer open to atmosphere Tap positions for a three-way tap (viewed from the side)


16

respirometer isolated



Measuring the rate of oxygen uptake

Answers 1 Simple answer: Oxygen molecules are absorbed by the organism and used in respiration. The same number of carbon dioxide molecules are released but these are absorbed by the soda lime. This reduces the pressure inside the test tube (fewer molecules 5 lower pressure). Atmospheric pressure pushes the liquid along the tube, until the pressure in and outside the tube is equal. Detailed respiration review answer: As above but should include reference to the role of oxygen as the final electron acceptor, and the fact that it eventually combines with hydrogen to make water. The carbon dioxide comes from the carbon dioxide released in the link reaction and the Krebs cycle as the carbohydrate is broken down. 2 a A drop in temperature inside the tube, or an increase in atmospheric pressure. b An increase in temperature inside the tube, or a decrease in atmospheric pressure. c The distance moved by the liquid in the control in each minute towards the organisms should be subtracted from the distance that the liquid in the experimental respirometer moved. Movement of the liquid in the control away from the organisms should be added to the distance in the experimental tube. 3 Diagram showing a very simple respirometer: Disadvantages – does not allow you to reset; it needs a control tube used alongside it; no scale so measurements likely to be less accurate. Advantages – very simple to set up; minimal number of connections makes a good seal easier to obtain. 1 cm3 syringe experimental tube screw clip three-way tap

small organisms

gauze KOH solution absorbs carbon dioxide

KOH solution manometer tube containing fluid

U-tube respirometer

Advantages – does not need to have an additional control as the second tube balances out the effects of changes in temperature or atmospheric pressure; the syringe allows you to move the liquid in the U to reset the apparatus. Disadvantages – tendency for the connections to leak in elderly school/college models (making the equipment useless); expense. 4 The experimental design should include either a U-tube respirometer or controls; a known mass of maggots; a range of temperatures (between 5 °C and 40 °C); a suitable increment between temperatures (say, 5 °C); repeated measurements (say, at least three at each temperature). Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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Technician 1 of 2

Practical 7.2 Investigating breathing

Purpose • To investigate lung volumes and rate of breathing.

Safety The safety guidelines in Sections 14.5 and 14.5 .1 in the CLEAPSS Laboratory Handbook should be followed when using a spirometer. This investigation is potentially hazardous. Students should be closely supervised at all times when using a spirometer. If the students are allowed to breathe through the spirometer for too long they can lose consciousness from lack of oxygen. Limiting the time spent breathing through the spirometer and carefully observing each student should prevent problems. When using oxygen and absorbing CO2: maximum time 5 minutes; in any other situation: maximum time 1 minute. If a student becomes less alert or has any feeling of suffocation they should stop immediately. Since the levels of CO2 are kept low by the soda lime, the students won’t be aware that they are running out of oxygen until it is potentially too late. A trained member of staff should use an oxygen cylinder to fill the spirometer. Use eye protection when handling soda lime. Soda lime is corrosive. Do not handle directly: use a spatula. Use a type of soda lime that changes colour when it is saturated, and replace it when it changes colour. Use 5–10 mesh particle size. Follow CLEAPSS guidelines on removing dust for spirometer use. A layer of polymer wool can be put at the inflow and outflow of the soda lime canister chamber to prevent dust getting into the chamber. Ensure that the soda lime canister is fitted so that air is breathed out through the canister. The subject will feel some resistance to breathing when using a spirometer. Because of this they should not use the spirometer while exercising. To investigate the effect of exercise, readings should be taken immediately after exercise. If resistance to breathing suddenly increases it may be due to valves in the spirometer sticking. If this occurs the valves may need to be replaced. Do not use oil, grease or glycerine on any part of the spirometer tubing as these may make explosive compounds with oxygen. Water with a little liquid detergent can be used to aid connection of tubes, etc. Keep all flames away from the spirometer. It is essential to follow good hygiene practice with regard to cleaning and disinfecting the mouthpiece and removing condensation and saliva from the tubes. Care should also be taken to choose the subject, avoiding any students with asthma or other breathing or circulatory problems. (Asthmatics may use a spirometer if they are otherwise in good health.) Read the manufacturer’s instructions carefully before using a spirometer.

In this activity students learn how a spirometer works and how to interpret the spirometer trace that is produced. Alternatively a datalogger can be used with a traditional water-filled spirometer or with an airflow spirometer. A range of different types of sensors can be used and some alternatives are described on the Teacher sheet. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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Investigating breathing

A spirometer can be used to investigate the effect of exercise on breathing rate and lung volumes. If you have not got access to a spirometer, measurement of vital capacity and tidal volume can be carried out using breath volume bags. These are a fraction of the cost of a spirometer. However, they cannot be used for measuring oxygen consumption. Requirements per student or group of students

Notes

Spirometer

If the spirometer has a pen it is worth checking that it is still working – they fail quite quickly. Always keep the ink reservoir with its piece of wire in place. Some new spirometers can be set up to feed the data into a computer. See notes on Teacher/ Lecturer sheet about using spirometers with dataloggers. Safety See notes on page 1.

Kymograph

See CLEAPSS Laboratory Handbook Section 15.1 for advice on speeds, etc. A chart recorder, computer or datalogger can be used in place of a kymograph.

Kymograph paper

Be aware that it matters which way up you attach the paper. If it is the right way up the pen moves smoothly over the join in the paper. If it is the wrong way up it catches on the join.

Soda lime for the spirometer canister

Safety See notes on page 1.

Disinfectant solution, ethanol or fresh 0.1% hypochlorite followed by rinsing with water

For rinsing mouthpiece and breathing tubes after use. To minimise an unpleasant taste on the mouthpiece, use Milton made up at the manufacturer’s dose. However, this takes 30 minutes to achieve disinfection.

Eye protection


156

Student 1 of 4


Purpose

Safety

• To investigate lung volumes and rate of breathing.

Use eye protection when handling soda lime. Soda lime is corrosive. Do not handle directly: use a spatula. A spirometer should only be used with supervision. If you have breathing or circulation (heart) problems or suffer from epilepsy you should not use the spirometer. Read the manufacturer’s instructions and safety notes before using the equipment. Stop using the spirometer at once if you experience any unusual breathing problems or feel dizzy or uncomfortable. (Asthmatics may use a spirometer if they are otherwise in good health.) A trained member of staff should use an oxygen cylinder to fill the spirometer.

Using a spirometer The apparatus shown below is a spirometer. Spirometers allow us to study both breathing and respiration. In this activity you will learn how a spirometer works and how to interpret the spirometer trace that is produced.

A spirometer

The general principle behind a spirometer is simple. It is effectively a tank of water with an air-filled chamber suspended in the water. It is set up so that adding air to the chamber makes the lid of the chamber rise in the water, and removing air makes it fall. Movements of the chamber are recorded using either a kymograph (pen writing on a rotating drum), a chart recorder, computer or datalogger.


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2 of 4 Student



Tubes run from the chamber to a mouthpiece and back again. Breathing in and out through the tubes makes the lid of the chamber fall and rise. The volume of air the person inhales and exhales can be calculated from the distance the lid moves. The apparatus can be calibrated so that the movement of the lid corresponds to a given volume. A canister containing soda lime is inserted between the mouthpiece and the floating chamber. This absorbs the CO2 that the subject exhales. In which direction will the pen move when the subject inhales?

Procedure You will need: • Spirometer • Kymograph, chart recorder or computer • Soda lime (for the spirometer canister)

• Disinfectant solution • Eye protection

Calibration In order to interpret the spirometer trace you need to know what both the vertical and the horizontal scales represent.

Finding the vertical scale The vertical scale measures the volume of air in the chamber. The spirometer’s floatingchamber lid has markings on it showing how much gas it contains. 1 First, empty the chamber completely and, if using a kymograph, make a mark on the paper while it is stationary, to show where the pen lies when there is no gas in the tank. Then force a known volume of air into the tank (e.g. 500 cm3) and make a second mark on the kymograph trace. 2 Repeat this procedure until the chamber has been completely filled with air. If using a kymograph, if the trace is too large or too small, the length of the arm supporting the pen can be adjusted so that the trace fits onto the paper. 3 Write the values next to your calibrating marks – they will help with interpretation of the trace later. Once the marks have been made on the paper it should be possible to count how many squares on the trace represent 1 dm3.

Finding the horizontal scale 4 On most kymographs there is a switch allowing you to set the speed at which the drum turns. Choose a speed close to 1 mm per second. This is your horizontal scale. Make a note of the speed on your trace, so that the trace can be interpreted once the experiment is complete.

Collecting data on breathing 5 After calibration, the spirometer is filled with oxygen. A disinfected mouthpiece is attached to the tube, with the tap positioned so that the mouthpiece is connected to the outside air. The subject to be tested puts a nose clip on, places the mouthpiece in their mouth and breathes the outside air until they are comfortable with breathing through the tube. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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Student 3 of 4



Volume 02/dm3

6 Switch on the recording apparatus and at the end of an exhaled breath turn the tap so that the mouthpiece is connected to the spirometer chamber. The trace will move down as the person breathes in. After breathing normally the subject should take as deep a breath as possible and then exhale as much air as possible before returning to normal breathing.

tidal volume

vital capacity

Time/minutes A sketch of a trace showing normal breathing and one forced breath in and out

A diagram of a spirometer trace is shown above. In this example the subject has breathed in and out normally three times, then taken as deep a breath in as possible, then forced the air from their lungs. Several pieces of information about the subject’s breathing can be read off this kind of trace, or worked out from it. • The tidal volume is the volume of air breathed in and out in one breath at rest. The tidal volume for most adults is only about 0.5 dm3. • Vital capacity is the maximum volume of air that can be breathed in or out of the lungs in one forced breath. • Breathing rate is the number of breaths taken per minute. • Minute ventilation is the volume of air breathed into (and out of) the lungs in one minute. Minute ventilation 5 tidal volume 3 rate of breathing (measured in number of breaths per minute). Some air (about 1 dm3) always remains in the lungs as residual air and cannot be breathed out. Residual air prevents the walls of the bronchioles and alveoli from sticking together. Any air breathed in mixes with this residual air.

Collecting data on rate of respiration Each time we take a breath, some oxygen is absorbed from the air in the lungs into our blood. An equal volume of carbon dioxide is released back into the lungs from the blood. When we use the spirometer, each returning breath passes through soda lime, which absorbs the carbon dioxide so, with the canister in place, less gas is breathed back into the spirometer Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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chamber than was breathed in. If we breathe into and out of the spirometer for (say) 1 minute, a steady fall in the spirometer trace can be seen. The gradient of the fall is a measure of the rate of oxygen absorption by the blood, and so is a measure of the rate of respiration by the body. 1 Using the trace produced in class, or one provided by your teacher/lecturer, find the following values: a tidal volume b vital capacity c breathing rate d minute ventilation. 2 Use the trace produced in class, or one provided by your teacher/lecturer, to work out the rate of oxygen consumption in someone at rest. 3 What differences would you expect if the subject had been exercising before a trace was taken? 4 Describe how you could use the apparatus to measure changes in breathing and respiration rates due to exercise. (Note that the apparatus you have used may not be suitable for use during exercise, and that measurements need to be taken immediately after exercise has stopped. Discuss with your teacher which is the best method for use with your apparatus.) State what exercise would be appropriate, and any hazards involved. Sketch the shape of the trace you would expect before and after exercise.


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Purpose • To investigate lung volumes and rate of breathing.

Spirometers A copy of a spirometer trace is provided at the end of these notes, for use when there is no access to a spirometer or as extra data to interpret. The practical describes the use of a spirometer with a kymograph. Alternatively a datalogger can be used with a traditional water-filled spirometer or with an airflow spirometer. A range of different types of sensors can be used and some alternatives are described below using a spirometer with a datalogger. If you have not got access to a spirometer, measurement of vital capacity and tidal volume can be carried out using breath volume bags. These are a fraction of the cost of a spirometer. However, they cannot be used for measuring oxygen consumption. Safety Use eye protection when handling soda lime. Soda lime is corrosive. Do not handle directly: use a spatula. A spirometer should only be used with supervision. Students with breathing or circulation (heart) problems or suffer from epilepsy should not use the spirometer. Read the manufacturer’s instructions and safety notes before using the equipment. CLEAPSS guidance is in section 14.5.1 of the Laboratory handbook. If the students are allowed to breathe through the spirometer for too long they can lose consciousness from lack of oxygen. Limiting the time spent breathing through the spirometer and carefully observing each student should prevent problems. When using oxygen and absorbing CO2: maximum time 5 minutes; in any other situation: maximum time 1 minute. If a student becomes less alert or has any feeling of suffocation they should stop immediately. Since the levels of CO2 are kept low by the soda lime, the students won’t be aware that they are running out of oxygen until it is potentially too late. Stop using the spirometer at once if the student experiences any unusual breathing problems or feels dizzy or uncomfortable. (Asthmatics may use a spirometer if they are otherwise in good health.) A trained member of staff should use an oxygen cylinder to fill the spirometer. If only a few breaths are to be measured, then atmospheric air is acceptable instead. The subject will feel some resistance to breathing when using a spirometer. Because of this they should not use the spirometer while exercising. To investigate the effect of exercise, readings should be taken immediately after exercise. If resistance to breathing suddenly increases it may be due to valves in the spirometer sticking. If this occurs the valves may need to be replaced. After use, the tubing from the spirometer should be removed and placed in 70% ethanol, to disinfect the internal ribbed surfaces where microorganisms might remain.

Notes on the procedure Effects of exercise on breathing rate and volumes can be investigated using the spirometer. Although this is not detailed on the activity sheet, students do need to be able to describe how to investigate the effects of exercise on tidal volume and breathing rate. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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Using a spirometer with a datalogger One advantage of collecting the data electronically is that the data can be moved from the datalogging software to a spreadsheet or a word processing program. The data collected can then be made available to students involved in the investigation. Sensors such as motion sensors can be used to record the up and down motion of the airfilled spirometer lid. A motion sensor can be mounted above the large top surface of the moving lid (see below). As the lid moves up and down the change in distance can be recorded. Make sure the limits of the motion sensor are known and the minimum recordable distance is available when the lid is at its closest to the sensor. If known series of gas volumes is introduced into the spirometer, and the distance between the motion sensor and the surface of the lid is recorded, it is possible to produce a calibration curve of distance versus volume. Most datalogging software can convert distance measurements recorded by the sensor to volume data. In a similar way, a linear motion sensor (e.g. rotary motion or pendulum sensor) can be attached to the spirometer as shown below. With a pendulum sensor, a string weighted with a small counterweight can be run from the end of the lid over the sensor’s pulley wheel. When the lid moves up and down, the string moves over the pulley. The sensor records a changing distance or angle. Calibration relating the volume of gas in the spirometer tank to the sensor’s reading will be required. rotary motion/position sensor A

B

pully

motion sensor

counterweight to tension cord cord

distance varies moving lid

moving lid

water

Carbon dioxide and oxygen levels in the spirometer lid can be datalogged. Monitoring these levels gives another safety check that the CO2 absorber is working and that O2 levels are not falling dangerously low. Alternatively, many of the datalogging companies now produce an airflow spirometer which measures volume by integrating the flow data over time. These devices are very lightweight, do not need carbon dioxide absorbers and do not need cylinders of medical oxygen (they use the oxygen from the atmosphere). Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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Answers 1 The values here are for the trace provided at the end of these notes. Obviously if a student’s own trace is used the answers may be different. a Tidal volume – between 0.5 and 0.8 dm3. Encourage students to take an average over several breaths. b Vital capacity 5 2.55 dm3 averaged over the two breaths. c Breathing rate 5 between 18 and 20 breaths per minute (depending on the section of the graph used). d Minute ventilation 5 19 breaths per minute × 0.65 dm3 5 12 dm3 min–1. 2 The rate of oxygen consumption determined from the trace at the end of these notes is approximately 1.0 dm3 min–1 for the first 30 seconds. 3 If the subject had been exercising, the rate of oxygen consumption would be higher, so the slope would be steeper. The trace would also show that the subject was breathing faster and more deeply. 4 This question allows students to think through the practicalities of using a spirometer in an investigation. They should think about the type of exercise: it is easier to use a spirometer immediately after running up and down stairs than after swimming or paragliding. They may also consider individual variation and sample sizes. They should also think about the limits imposed by the amount of oxygen available in the tank. Comments on the effect of the spirometer on the subject are also worth encouraging – a spirometer often causes breathing to become slightly more laboured particularly with older models that have smaller diameter tubing. The graph should show that breaths become deeper and more frequent after exercise, with a greater rate of oxygen consumption. After exercise

Volume 02/dm3

Before exercise

Time/minutes


20




1 cm

Spirometer trace for spirometer used with soda lime and filled to 9 dm3 with oxygen

9

Volume 02(dm3)

8

7

6

5 Kymograph speed: 64 mm min�1

Sample reference spirometer trace


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Technician 1 of 1

Practical 8.1 Can snails become habituated to a stimulus?

Purpose • To investigate habituation of snails to a stimulus.

Requirements per student or group of students

Notes

Giant African land snail (or large garden snail)

Giant African land snails can be purchased in some pet shops or from Biological suppliers such as Blades Biological. If purchasing giant African land snails they should be bred in this country, to reduce chance of them having parasites. They are easy to keep in a glass or plastic aquarium tank containing several centimetres of compost, organic compost is recommended. There are several publications available about the care of land snails. If giant land snails are not available, a large garden snail can be used. Snails should be kept in moist conditions before the lesson, to encourage activity. If snails are still inactive, they can be paced for a short time in warm water. Safety Wash your hands thoroughly after handling the snail. Equipment and surfaces should be disinfected with 1% Virkon.

Cotton wool bud Small beaker

Any small container for water to allow cotton wool bud to be dampened.

Board

Clean, firm surface for the snails, for example a plastic chopping board.

Stop watch


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Student 1 of 2


Purpose • To investigate habituation of snails to a stimulus.

Touching snails Lots of people, at some time in their childhood, will have touched a snail in the garden and noticed that it withdraws its eye stalks into its body. For such a slow-moving animal this seems a very quick response, this suggests it is an important response for protection and survival. A snail only withdraws into its shell when it is either inactive or threatened. When touched, it withdraws to avoid danger. Do snails become habituated to the stimulus, ceasing to withdraw with repeated stimulation? In this investigation you will collect data to find out if habituation to a touch stimulus does occur in these organisms. Safety Wash your hands thoroughly after touching the snails once all the equipment has been put ready for disinfection. Take care that the stimulus causes no harm to the snails.

Procedure You will need: • One giant African land snail (or a garden snail if not available) • One dampened cotton wool bud • Suitable clean, firm surface for the snails (e.g. a plastic chopping board) • Stopwatch

1 Collect one giant African land snail, and place it on a clean, firm surface. Allow the snail to get used to its new surroundings for a few minutes until it has fully emerged from its shell. 2 Dampen a cotton wool bud with water. 3 Firmly touch the snail between the eye stalks with the dampened cotton wool bud and immediately start the stopwatch. Measure the length of time between the touch and the snail being fully emerged from its shell once again, with its eye stalks fully extended. 4 Repeat the procedure in step 3 for a total of 10 touches, timing how long the snail takes to re-emerge each time. 5 Record your results in a suitable table. 6 Present your results in an appropriate graph. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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Can snails become habituated to a stimulus?

Questions 1 Write a hypothesis which this experiment will test. 2 Using your graph, state if you think there is a positive, negative, or no, correlation between the number of stimulations and the time for eye stalk withdrawal. 3 Explain any patterns or trends in your data, supporting your ideas with evidence from the data and your biological knowledge of habituation. Relate your findings to your hypothesis. 4 Suggest a reason why snails may become habituated to a prodding stimulus in the wild. 5 Evaluate the procedure used for this experiment. 6 This experiment has been shown to be less successful if the snails are handled regularly prior to the experiment. Suggest why handling prior to the experiment could affect the results of the experiment.

Going further 7 Write a null hypothesis that this experiment will test. 8 Complete a Spearman’s rank rs correlation test to determine if there is a statistically significant correlation between the variables. A table with the headings below will help. Number of times the snail has been stimulated

Rank stimulation

Time/seconds

Rank time

Difference/D

D2

9 Use a table of critical values to accept or reject your null hypothesis. If your calculated Spearman rank value (rs) is greater than the critical value, then the null hypothesis is rejected. If your calculated rs value is less than the critical value, then the null hypothesis is accepted. 10 Write a statistical conclusion for your experimental data. Make sure you include: • your calculated value of rs • the number of pairs of data • the significance level • the critical value • whether the null hypothesis is being accepted or rejected.


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Purpose

Safety

• To investigate habituation of snails to a stimulus.

Wash your hands thoroughly after touching the snails once all the equipment has been put ready for disinfection. Take care that the stimulus causes no harm to the snails.

Notes on the procedure In this investigation students find out if habituation to a touch stimulus occurs in snails. A giant African land snail is best, but large garden snails can be used. The student sheet that accompanies this activity contains a procedure for this experiment but students could be asked to plan the investigation themselves. The experiment is more reliable if snails are handled little prior to the experiment, so avoiding them becoming habituated to stimuli. The completion of Spearman’s rank correlation provides an extension to this activity. The null hypothesis to be tested would be: there will be no correlation between the number of stimuli the snail receives and the time taken for the snail to re-emerge. Further investigations might include how long the habituation lasts, or how handling prior to the experiments affects the results.

Answers 1 As the number of stimuli increase, the time taken for the snail to re-emerge will decrease – a negative correlation. 2 Students present the results as a scatter graph, with the number of stimuli on the x-axis, and the time taken to re-emerge on the y-axis. Students should make a simple statement, from their results, as to whether there appears to be a positive, a negative, or no, correlation. 3 There is a negative correlation – as the number of stimuli increase the time taken for the snail to re-emerge decreases. Students should make a reference to the data. With repeated stimulation, Ca21 channels in the presynaptic membrane become less responsive. Less Ca21 crosses the membrane into the presynaptic (sensory) neurone. As a result less neurotransmitter is released into the synaptic cleft. This means that an action potential across the postsynaptic membrane is less likely. Fewer action potentials are produced in the postsynaptic motor neurone so less of a response is observed. 4 The snail learns that the stimulus is not causing harm, and so it is withdrawing unnecessarily. This effect may be used in its natural habitat when faced with repetitive stimuli, such as vegetation touching the head/eye stalks. 5 Relevant comments would include: the need for replication using snails of approximately the same size and age; the need to control of the size and position of the stimulation; the need to control other variables that may affect the response, e.g. drying out of the snail; difficulties in determining when the snail has fully extended (measuring the eye stalk length prior to stimulation may overcome this problem); effect of handling the snail prior to the experiment. 6 If the snails have been handled too much prior to the experiment, they may have already become habituated to this type of stimulus so further stimulations will not change the response. Edexcel practical materials created by Salters-Nuffield Advanced Biology, ©University of York Science Education Group.

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Edexcel A2 Biology Practicals (Complete)

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