Edexcel A2 Biology Practicals (Complete)

8/17/2019 Edexcel A2 Biology Practicals (Complete)

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

Technician 1 of 1

Practical 5.1 Looking for patterns and correlations

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

• To carry out a study on the ecology of

a habitat.

Purpose

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.

Safety


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Student 1 of 8

Practical 5.1 Looking for patterns

Observing patternsHave 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 patternsLook 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 – thestudents 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.

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

Purpose


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2 of 8 Student

Practical 5.1 (cont.) Looking for patterns

Completing a transect studyOne 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.

Procedure1 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 eldwork.

A pilot study in advance of the main data collection will help you make these decisions.

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.

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

peg marked ‘20’

origin

peg marked‘0’

21 22 23 24 25 26 27 28 29 30

0.5 m� 0.5 m

quadratno. 6

transectcontinues

tape case


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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 attwo 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

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 randomnumbers 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.

Figure 1 Using measuring tapes to define a sample area.

1

2

3

4

5

1 2 3 4 5

position of 0.25 m2

quadrat using randomnumbers 2, 4

Tape measure 1

T a p e m e a s u

r e 2


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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 ofPleurococcus

(analga) 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 samplingRandom 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:

% cover 5 hits

____________

hits1 misses 3 100


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6 of 8 Student


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.

multiple hit

metal spike(such as atent peg)inserted inground

holesknitting needle


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Student 7 of 8


Figure 3 Net and traps for sampling animals.

• Pitfall trap for sampling arthropods• Tullgren funnel for collecting organisms from the soil or leaf litter

• Pooter for collecting insects

• Sweep net

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

flat stone preventsrainfall filling trap

stick supportsoil sample

25 W bulb

60 W bulb

rod for supporting bag

water

rubber tubing clip

beaker

glass funnel

soil sample inmuslin bag

16 mesh flour sieveground slopesaway from trapfor drainage

bait of meator ripe fruit

glass collecting tube

clear plastictube

glass mouthpiece 80% alcohol

polythenefunnel

Organisms moveaway from the heatand light, fallinginto the jar.

gauze coveringtube opening

air suckedthroughmouthpiece

air and arthropodsdrawn into tube

cork or rubber bung

specimen tubewhere arthropodscollect

jam jar sunk intosoil

This net is swept throughlow-growing vegetation,collecting any animalsin the mesh net.


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8 of 8 Student

Measuring abiotic factors when sampling the environmentAngle 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.

LightUse 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:

% soil moisture 5 mass of fresh soil 2 mass of dry soil

________________________________

mass of fresh soil 3 100

Soil organic matterA 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:

% organic matter in soil 5 mass of dry soil 2 mass of burnt soil

________________________________

mass of dry soil 3 100

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 cm

3

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.



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Teacher/Lecturer 1 of 3

Practical 5.1 Looking for patterns

Notes on the procedureStudents 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 onlineresources 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 carefullyThe 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.

• To carry out a study on the ecology of

a habitat.

Purpose

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.

Safety


11/458


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-owing 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 identication

• 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.


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Class organisationStudents 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 resourceFieldWorks 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

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.

• 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.

Purpose


Brine shrimp egg cysts Available from pet shops. There are approximately 24 000 egg cystsper gram so only tiny quantities are required. Brine shrimps willbreed and produce cysts that can be collected. They adhere to thesides 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.

40 cm3 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 eachtemperature to be investigated)

A range of temperatures between 5 and 35 °C is recommended. Theoptimum 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 secondday.

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 andalgae, set up well in advance. Details in the Dockery andTomkins book

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


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Student 1 of 2


brine shrimps

Brine shrimpsBrine 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.

Drawings to show features of brine shrimps

Procedure

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 cm

3

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.

• To investigate the effect of temperature on the hatching success of brine shrimps.




Purpose

1

mm

eggs

secondantenna

female(brownish red)

firstantenna

male(blue/green)

• 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

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)


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2 of 2 Student

Practical 5.2 (cont.) 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 faras 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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brine shrimps

Notes on the procedure

If students are given basic information on maintaining brine shrimps this activity could beplanned 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.

• To investigate the effect of temperature on the hatching success of brine shrimp.




Purpose


17/4533


Student 1 of 1

Practical 6.1 DNA gel electrophoresis

Procedure

You may have the opportunity to complete experimental work using restriction enzymes and

gel electrophoresis or you may use the simulation of this.

• To use gel electrophoresis to separate DNA

fragments of different sizes.

Purpose

If you undertake gel electrophoresis make sure

you are aware of the hazards and follow the

instructions of your teacher very carefully.

Safety


18/4511



Practical 6.1 DNA gel electrophoresis

Restriction enzymesThe use of restriction enzymes to cut DNA and electrophoresis to separate the resulting

fragments is possible using equipment available from NCBE (National Centre forBiotechnology 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 usingrestriction 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.

• To use gel electrophoresis to separate DNA

fragments of different sizes.

Purpose

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.

Safety


19/4512



Practical 6.1 (cont.) DNA gel electrophoresis

Pfizer and Bio-Rad have donated biotechnology equipment to 31 ITT centres around thecountry 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 Bio-

Rad 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.


20/4534


1 of 1 Student

Practical 6.2 DNA amplification using PCR

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). Comparingthe 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.

• To use PCR to amplify DNA.

Purpose

If you undertake PCR make sure you are aware

of the hazards and follow the instructions of

your teacher very carefully.

Safety


21/4513



Practical 6.2 DNA amplification using PCR

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 teachersand 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!

• To use PCR to amplify DNA.

Purpose

If PCR is to be used a full risk assessment

should be obtained for the procedure and

carefully followed by both staff and students.

Safety


22/4514



Practical 6.2 (cont.) DNA amplification using PCR

Contact details and examples of the protocols offered by NCBE, Bio-Rad and EdvotekEurope:

Organisation Example of suitable protocols

NCBENational Centre for Biotechnology Education

Science and Technology CentreThe University of ReadingWhiteknightsReading,RG6 6BZwww.ncbe.reading.ac.uk

0118 987 [email protected]

Amplifying lambda DNA (from the Illuminating

DNA booklet)Investigating plant evolutionAnalysing mitochondrial DNA. This practicalrequires a microcentrifuge and the protocol is notdownloadable from the NCBE website. It is basedon a protocol developed for a DNA workshop at theCold Spring Harbor Laboratory. It can be obtained

direct from NCBE.

Bio-RadBio-Rad Laboratories Ltd.Bio-Rad HouseMaxted RoadHemel HempsteadHertfordshireHP2 7DX

www.biorad.com/ Follow the links toLife Science education, then About BiotechnologyExplorerFreephone: 0800 181134

Crime scene investigator – PCR basicsPV92 PCR infomatics kitGMO investigator kit (only useful if GM foodstuffsare easily available)

Edvotek EuropeThe Biotechnology Education CompanyPO Box 280Hertford

SG13 9DGwww.edvotek.co.ukFollow links to Experiments and then Polymerasechain [email protected]

332 Mitochondrial DNA analysis using PCR


23/45153

Technician 1 of 1


Practical 6.3 Which antibiotic is most effective?

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.

• To investigate the effect of different antibiotics on bacteria.

Purpose


Agar plates seeded with bacteria or equipment for their

preparation (i.e. access to one of two broth cultures of knownbacteria; Universal McCartney bottle of nutrient agar; sterilePetri dish; sterile pipette)

Suitable bacteria are listed in various catalogues. Phillip Harrisuse E. coli (K12 strain) and Staphylococcus albus in their kit.The Student sheet with instructions for preparation of seededagar plates, ‘Pouring agar plates’, can be found in Edexcel ASBiology 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

Forceps (metal)Autoclaved after placing in cotton wool stoppered boilingtube.

A Mast ring or separate antibiotic discs Separate antibiotic discs are usually cheaper, but it isimpossible 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.


24/4535



• To investigate the effect of different

antibiotics on bacteria.

Purpose

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.

Safety

IntroductionWhen 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

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.

• Marker pen

• Autoclaved forceps

• Mast ring or antibiotic-impregnated paper

discs

• Adhesive tape

• Eye protection

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

Student 1 of 2


25/4536


Practical 6.3 (cont.) Which antibiotic is most effective?

5 Wash your hands with soap or handwash and clean the bench again using the Virkonspray.

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?

2 of 2 Student


26/4515




Notes on the procedureThe 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 tapefrom 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.

Answers1 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 notkill the bacteria. Some antibiotics can be used together and produce a larger effect when

combined than if administered separately. This is known as synergism.

• To investigate the effect of

different antibiotics on bacteria.

Purpose

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.

Safety


27/45154

1 of 1 Technician


Practical 7.1 Measuring the rate of oxygen uptake

RespirometersA 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, orpossibly the organisms are too cold, or dead!

• To demonstrate the uptake of oxygen in respiration.

• To measure the rate at which an organism respires.

Purpose


Respirometer See the diagram on the Student sheet. If a pipette is used,the scale shown will not be needed. Ideally the syringes areattached with a three-way tap. If these are not available a

rubber tube and clip can be used. U-tube respirometers wouldbe even better, if there is a class set available. See also section15.10 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, ormaggots, 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 inuse) 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


28/4537



Respirometers

Respirometers range from relatively simple pieces of equipment used in school sciencelabs 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

1 Assemble the apparatus as shown in the diagram below.

A simple respirometer

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

• To demonstrate the uptake of oxygen in

respiration.

• To measure the rate at which an organism

respires.

Purpose

Wear eye protection when handling soda lime.

Soda lime is corrosive. Do not handle directly:

use a spatula.

Safety

scale

1

cm3 pipette or glass tube

syringe

glass tubing

small organisms

gauze

soda lime

three-way tap

coloured liquid

• Soda lime

• Coloured liquid• Dropping pipette

• Permanent OHT marker pen

• Solvent (to remove the marker)

• Cotton wool• Stopclock

• Eye protection

You will need:

• Respirometer (see diagram

below)• 5 g of an actively respiring

organism

Student 1 of 2


29/4538


Practical 7.1 (cont.) 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 p r 2 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 oneyou are using?

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.

sodalime

wiremesh

organism tobe studied

capillarytube

drop of liquid

2 of 2 Student


30/4516


Notes on the procedure

Rate of respiration is not a learning outcome in the specification. The respirometer shown inthe 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 p r 2 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.

Tap positions for a three-way tap (viewed from the side)

• To demonstrate the uptake of oxygen in

respiration.

• To measure the rate at which an organism

respires.

Purpose

Wear eye protection when handling soda lime.

Soda lime is corrosive. Do not handle directly:

use a spatula.

Safety

respirometer opento syringe

respirometer opento atmosphere

respirometer isolated

31 2




31/4517


Practical 7.1 (cont.) Measuring the rate of oxygen uptake

Answers1 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.

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).

KOH solutionabsorbs carbondioxide

KOH solution

three-way tap

1

cm3 syringe

gauze

screw clip

experimental tube

manometer tubecontaining fluid

small organisms



32/45155


Practical 7.2 Investigating breathing

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-filledspirometer or with an airflow spirometer. A range of different types of sensors can be used

and some alternatives are described on the Teacher sheet.

• To investigate lung volumes and rate of breathing.

Purpose

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 replaceit 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 aidconnection 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.

Safety

Technician 1 of 2


33/45156


2 of 2 Technician

Practical 7.2 (cont.) Investigating breathing

A spirometer can be used to investigate the effect of exercise on breathing rate and lungvolumes. 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.


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


34/4539


Practical 7.2 Investigating breathing

Using a spirometerThe 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), achart recorder, computer or datalogger.

• To investigate lung volumes

and rate of breathing.

Purpose

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 goodhealth.)

A trained member of staff should use an oxygen cylinder to fill

the spirometer.

Safety

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Tubes run from the chamber to a mouthpiece and back again. Breathing in and out throughthe 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

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 floating-

chamber 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 breathing5 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.

• Disinfectant solution

• Eye protection

You will need:• Spirometer

• Kymograph, chart recorder or computer

• Soda lime (for the spirometer canister)

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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.

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 inand 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 ventilation5 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 respirationEach 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

vitalcapacity

tidalvolume

V o l u m e 0 2 / d m 3

Time/minutes

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chamber than was breathed in. If we breathe into and out of the spirometer for (say) 1minute, 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

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Edexcel A2 Biology Practicals (Complete)