A level practicals

1 Remove 1 Daphnia and place in cavity slide. Remove pond water and replace with distilled water. 

2 Leave for 5mins to acclimatise then count heart rate under microscope for 30s, multiply number by 2 to calculate beats/min.

3 Repeat with 2 more Daphnia. 

4 Repeat again, this time with small concentration of caffeine solution in place of distilled water. 

5 Carry out for 5 concentrations of caffeine 

1 Pipette 1 cm3 of 1% DCPIP solution into a test tube.

2 Record the start volume of 1% vitamin C solution in a burette. Add it slowly to the DCPIP solution until the blue colour of the DCPIP has just disappeared. Record the end volume. 

3 Repeat this procedure with the fruit juices provided. If only one or two drops of the fruit juice decolourises the DCPIP, dilute the juice and repeat the test.

4 The 1% vitamin C solution contains 10 mg of vitamin C in 1.0 cm3. Calculate the mass of vitamin C that is required to decolourise 1 cm3 of the DCPIP solution. Use this value to work out how much vitamin C each of the fruit juices contain, in mg cm–3. 

1 Cut cylindrical samples from a single beetroot. Cut eight 1 cm length sections from these samples. 

2 Place one of the beetroot sections into a boiling tube containing 5 cm3 distilled water. This is 0% alcohol concentration. Repeat with seven test tubes containing 10%- 70% alcohol. 

3 Remove beetroot sections using a technique that does not squeeze the slice. 

5 Switch on the colorimeter and set it to read percentage absorbance. Set the filter dial to the blue/green filter.

6 Adjust the colorimeter to read 0 absorbance for clear water. Don't alter the setting again during the experiment.

7 Place 2 cm3 of the dye solution into a colorimeter cuvette and take a reading for absorbency. Repeat the readings for all the alcohol concentrations.

1 Cut cylindrical samples from a single beetroot. Cut eight 1 cm length sections from these samples. 

2 Place eight labelled boiling tubes, each containing 5 cm3 distilled water, into water baths at 0-70 C. Leave for 5 minutes until the water reaches the required temperature. Place one of the beetroot sections into each of the boiling tubes. Leave for 30 minutes in the water baths.

3 Remove beetroot sections using a technique that does not squeeze the slice. Shake the water/solution to disperse the dye.

5 Switch on the colorimeter and set it to read percentage absorbance.  Set the filter dial to the blue/green filter.

6 Adjust the colorimeter to read 0 absorbance for clear water. Do not alter the setting again during the experiment.

7 Place 2 cm3 of the dye solution into a colorimeter cuvette and take a reading for absorbency.Repeat the readings for all the temperatures. 

1) Measure 5, 10, 15 and 20 cm³ of trypsin into four separate beakers. Then add enough water to make it up to 20 cm³. Also, measure out 20 cm³ of water in another beaker. This will mean you will have six solutions: 0% trypsin, 25% trypsin, 50% trypsin, 75% trypsin and 100% trypsin.

2) Measure 50cm3 of milk powder solution. Then measure the temperature. 

3) Put the pH9 buffer into a beaker.

4) Add the 100% trypsin solution to 50cm3 of milk powder solution to the same beaker.

5) Immediately start timing the reaction, measuring the temperature every 10 seconds for five minutes.

6) Once the five minutes is over, wash the beaker out thoroughly and repeat the experiment five times for the 100% solution.

7) After repeating the 100% solution five times, repeat the method five times each for the other concentrations of trypsin.

1 Cut off about 5 mm from several root tips of some growing garlic roots using fine scissors. Choose root tips that are white and have a firm, rounded end.

2 Put the root tips into a hollow glass block or small sample tube containing 2 cm3 1 M HCl for exactly 5 minutes.

3 Put the root tips in a watch glass containing approximately 5 cm3 cold water. Leave the root tips for 4–5 minutes, then dry them on filter paper. Take care – the root tips will be very fragile.

4 Transfer one of the root tips to a clean microscope slide.

5 Gently break up the root tip with a mounted needle  Add one small drop of toluidine blue and leave to stain for 2 minutes.

6 Cover with a coverslip and blot firmly with several layers of tissue or filter paper. Press gently to spread the root tip, or tap gently on the coverslip with the end of a pencil.

7 View under the microscope and look for cells with visible chromosomes. If cells are overlapping, squash the slide again between two wads of filter paper. 

1 Place a small piece of tinned rhubarb on a watch glass. Use forceps to pick out one or two vascular bundles from this block of tissue and place them on a microscope slide.

2 Use mounted needles to tease the vascular bundles apart. Cover the tissue with a drop of methylene blue, and leave for 5 minutes.

3 Draw off the extra stain with filter paper. Place a drop of dilute glycerol on the fibres and mount under a coverslip.

4 Examine your preparation under a microscope. If the tissues are not separated enough, place your slide on a piece of filter paper, put a filter paper pad on the coverslip and press down with your thumb. This may separate out the tissue. Do not move your coverslip sideways at all. You may need to re-irrigate the slide with glycerol after squashing it. To do this, place a drop of glycerol on the slide next to the coverslip. It will be drawn under the coverslip by capillary action. Blot off any excess and re-examine the slide.

5 Look for vascular bundles amongst the separated tissues 

1 Half fill a tube with the ‘all nutrients present’ solution. 2 Cover the top of the tube with foil or paraffin and push down on covering so that there is a well in the centre. Gently push the geranium stem/roots of Mexican hat plantlet through the hole so it is in solution below.

3 Repeat with solutions lacking in nitrogen or phosphate or potassium or magnesium or calcium or lacking all. 

4 Wrap all tubes in aluminium foil and place in tube holder on sunny window sill. Observe regularly

1 Plant material should be left to soak in a bucket of water for about a week in order for the fibres to be easily extracted

2 Once fibres removed, connect between 2 clamp stands and gradually add mass in the middle until the fibre snaps. 

3 Try with individual fibres from different plants and different ways of combining fibres eg twists and plaits. 

1 Agar plates seeded with suitable bacteria need to be prepared. 

2 Obtain a plant extract by crushing 3 g of plant material with 10 cm3 of industrial methylated spirit and shake it from time to time for 10 minutes. Pipette 0.1 cm3 of extract onto a sterile antibiotic assay paper disc. 

3 Repeat steps 1 and 2 for other plants, making separate test discs for each extract.

4 Use sterile forceps to place the test discs onto the bacterial plate together with the suitable control per plate. Ensure that you can distinguish between the different discs

5 Close the Petri dish and tape it. Do not tape all round the dish because this can lead to the growth of anaerobic bacteria, some of which may be harmful. 

6 Incubate the plates for 24 hours at 25 °C. Observe the plates without opening them. Bacterial growth on an agar plate looks cloudy. Make any appropriate measurements that will enable you to compare the antibacterial properties of the different plant extracts.

7 Wash your hands thoroughly with soap and water after completing the practical.

Random sampling = set up grid using tape measure, use random numbers to generate points to place quadrat to collect data.

Systemic sampling = line transect often used especially to study zonation. A tape measure is laid along several zones to be looked at and quadrats are used to record data at regular intervals

Measuring abundance: Density = presence of organisms per quadrat

Frequency = percentage of quadrat squares containing organism

Percentage cover = percentage of ground covered with organism in a quadrat 

Pitfall trap = to collect invertebrates

Pooter = to collect invertebrates into a container

Tullgren funnel = to collect organisms from soil or leaf litter

Baermann funnel = to collect living organisms from water  

1 Cut three small green spinach leaves into small pieces with scissors, but discard the tough midribs and leaf stalks. Place in a blender containing 20 cm3 of cold isolation medium 

2 Blend for about 10 s.

3 Place four layers of muslin or nylon in a funnel and wet with cold isolation medium.

4 Filter the mixture through the funnel into the beaker and pour the filtrate into pre-cooled centrifuge tubes supported in an ice-water-salt bath. Gather the edges of the muslin, wring thoroughly into the beaker and add filtrate to the tubes.

5 Check that each centrifuge tube contains about the same volume of filtrate.

6 Centrifuge the tubes for about 10 mins to get a small pellet of chloroplasts

7 Pour off the supernatant into a boiling tube being careful not to lose the pellet. Resuspend the pellet with about 2 cm3 of isolation medium, using a glass rod. 

8 Put DCPIP into the solution and put in front of a lamp. It should change colour

1 Puree some frozen peas, with some water, in a blender. Filter this to get just the pea juice

2 Pour 10cm3 of 20% volume hydrogen peroxide into a test tube. 

3 Put the test tube into a beaker filled with water at room temperature.

4 Set up a bung with a delivery tube to a gas collection syringe. 

5 Using a syringe, put 5cm3 of the pea filtrate into the test tube with the hydrogen peroxide. 

6 Make a note of the volume of oxygen collected every 20 seconds for two minutes.

7 Repeat the experiment, using ice, water at 30oC, and water at 40oC. At each temperature, do the experiment at least three times. 

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 and then add 100 cm3 of deionised water and stir until the salt completely dissolves.

4 Label the beaker with the temperature at which it will be incubated.

5 Place a tiny pinch of egg cysts onto a large sheet of white paper.

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. 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 Incubate the beakers at the appropriate temperatures. 

9 Record the number of larvae that have successfully hatched at each temperature.  

1 Mix DNA with desired restriction enzyme and loading dye. 

2 Prepare agar and pour into electrophoresis mould. 

3 Once set, fill electrophoresis tank with buffer solution. Use micropipette to load restriction ladder into first well then DNA samples cut with restriction enzyme into the other wells. 

4 Connect to electrical supply, turn on and leave until the dye has moved to the opposite end of the gel tank.

5 Switch off and disconnect electrical supply. Carefully remove the gel from the tank and view under UV light. 

1 For this practical you will need to work in sterile conditions (aseptic technique) 

2 Prepare an agar plate seeded with bacteria. 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 a Mast ring. Raise the lid of the Petri dish and place the Mast ring firmly in the centre of the agar;

4 Tape the dish securely with two pieces of adhesive tape (but do not seal it completely), then keep it upside down at 30°C for 48 hours. 

5 After incubation, look carefully at the plate but do not open it.

6 Measure the diameter of the inhibition zones in millimetres

1 Assemble the apparatus as shown in Figure 2 of activity 7.7.

2 Place 5 g of maggots into the boiling tube and replace the bung. 

3 Introduce a drop of marker fluid into the pipette using a dropping pipette. Open the three-way tap to the syringe and move the fluid to a convenient place on the pipette if needed

4 Mark the starting position of the fluid on the pipette with a fine permanent 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 after 5 minutes.

6 At the end of 5 minutes open the connection to the outside air.

7 Measure the distance travelled by the liquid after 5 minutes

8 If your tube does not have volumes marked onto it you will need to convert the distance moved into volume of oxygen used.

1 First, empty the chamber completely and make a mark on the kymograph paper to show where the pen lies when there is no gas in the tank. Then force a known volume of air into the tank and make a second mark on the kymograph trace. Repeat this procedure until the chamber has been completely filled with air. 

2 Write the values next to your calibrating marks – they will help with interpretation of the trace.

3 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. Make a note of the speed on your trace.

4 After calibration, the spirometer is filled with medical grade 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.

5 Switch on the kymograph. At the end of an exhaled breath turn the tap so that the mouthpiece is connected to the spirometer chamber. 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

1 Collect one giant African land snail and place it on a clean, firm surface. Wait for a few minutes until the snail has fully emerged from its shell and is used to its new surroundings.

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 ten touches, timing how long the snail takes to reemerge each time.