Technician otes Biology - Pearson Education...Practical 3: Production of a dilution series of a solute to produce a calibration curve with which to identify the water potential of

Technician Notes

1 © Pearson Education Ltd 2018

BiologyAQA AS and A Level

Practical 1: Investigation into the effect of a named variable on the rate of an enzyme-controlled reaction

Objectives

● To be able to measure the initial rate of enzyme activity

● To understand why measuring the initial rate is important

Safety ! Equipment

● Trypsin solution at concentrations of 1% or above is an irritant.

● Wash splashes from the skin as quickly as possible.

● Wear eye protection.

● skimmed milk powder suspension (2%)

● standard protease (trypsin) solution (1%)

● test tubes and holder

● pipettes

● eye protection

● access to colorimeter (see diagram) (or light meter with data logger)

● two cuvettes

● distilled water

Background

In this practical students investigate the breakdown of milk protein (casein) by protease enzymes such as trypsin. The reaction is monitored using a colorimeter.

Procedure

1. The 1% trypsin solution is diluted with distilled water to produce test solutions of 0.2%, 0.4%, 0.6% and 0.8%. The aim is to produce 10 cm3 of each concentration.

2. Students place 2 cm3 of trypsin solution and 2 cm3 of distilled water in a cuvette, to use as a reference cuvette when setting the colorimeter absorbance to zero.

3. 2 cm3 of milk suspension is then measured into a second cuvette.

4. Students then add 2 cm3 of trypsin solution to the milk in the cuvette. Working quickly, they mix the trypsin solution and milk, place the solution in the colorimeter and start the data logger.

5. The absorbance should be measured immediately and then at 15-second intervals (or more frequently if students are recording electronically) for 5 minutes, or until there is little change in absorbance.

6. The cuvette should be rinsed with distilled water and the procedure repeated for each concentration.

absorbancereadouttransmitted light

light detectorcuvettecoloured filter

light source

solutionabsorbs

light

BiologyAQA AS and A LevelTechnician Notes


Objectives

● To know how to prepare a temporary slide of a root tip to observe mitosis

● To recognise the stages of mitosis in dividing cells

● To identify hazards, associated risks and control measures for the procedure

● To use an eyepiece graticule

● To calculate magnification

Safety ! Equipment

● Eye protection must be worn.

● Take care with glassware and scissors.

● Acetic orcein stain is corrosive, causes burns, has an irritating vapour and will stain. Wear eye protection and avoid contact with skin. If contact does occur, wash the area thoroughly with water for 10 minutes. Mop up spillages immediately.

(per student or pair of students)

● garlic clove with growing root tip

● glass slide and coverslip

● scissors

● water bath at 55 °C

● small bottle with lid or laboratory stretch film

● hydrochloric acid 1 mol dm−3

● acetic orcein stain in a small bottle or vial

● two dissecting needles

● paper towels

● optical microscope

● eyepiece graticule

● white tile

● fine forceps

● stop clock

● safety information sheet

Background

In this practical, students prepare stained squashes of cells from plant root tips to identify the stages of mitosis. Hydrochloric acid is used to break down the pectins which hold the plant cells together; the preparation is then treated with acetic orcein to stain the chromosomes dark red and to stop mitosis by fixing the cells.

Students examine their preparations using a microscope, and produce labelled pictures of the cells during mitosis.

Procedure

Part 1: Mitotic indexEnsure that students follow all safety precautions stated above, and that they complete a risk assessment before beginning this practical.

1. In this practical, students are asked to prepare 1 mol dm–3 hydrochloric acid solution. You may wish to prepare this for the students ahead of the lesson and decant it into the small bottles, depending on the exact protocol followed. Students then place the bottles in the thermostatically controlled water bath at 55 °C, and leave them for 15 minutes.

2. Students place a garlic clove in the top of the bottle so the roots are submerged in the hydrochloric acid at 55 °C. They must leave roots in the acid for 5 minutes.

Practical 2: Preparation of stained squashes of cells from plant root tips; set-up and use of an optical microscope to identify the stages of mitosis in these stained squashes and calculation of a mitotic index

Technician Notes



Procedure continued

3. After 5 minutes, the clove is removed from the bottle and the roots rinsed thoroughly in tap water. Students use a pair of sharp scissors to cut off several root tips 5–10 mm in length. The root tips are dropped into a small vial of acetic orcein standing on a white tile, making sure the root tips are immersed in the stain. A lid or laboratory stretch film is then placed over the vial. This lid should have a pin-prick hole, or be slightly loose if it is a screw cap, to prevent the ejection of liquid during heating.

4. Students place the vial containing the root tips in acetic orcein in the 55 °C water bath for 5 minutes to intensify the staining.

5. After 5 minutes, forceps are used to take the tips out of the vial and place them on a microscope slide. A drop of water is added to the root tip on the slide. The root tips are teased apart with needles (maceration), to spread out the cells a little, and covered with a coverslip. Students are then asked to replace the lid on the vial of stain and return it to the teacher.

6. The slide is wrapped in several layers of paper towel and the paper gently pressed to squash the tissues. Students should take care not to twist the slide as they press down on it, or the coverslip will break.

7. Students examine the slide under the microscope on low power to identify the area of dividing cells or meristem (see Figure A), positioning the cells in the centre of the field of view. Meristem cells are small and square, have no obvious vacuoles and are usually found in rows.

meristem

Figure A: Section of a root tip showing meristem with dividing cells

8. With the microscope power set high (×400), students identify as many stages of the cell cycle in their field of view as possible.

9. They count number of cells in each of the stages of mitosis, plus interphase, in the field of view and record their results in a table.

10. Students are asked to draw and annotate one cell from each of the stages they have identified. Their drawings should be simple outlines of the cells and the groups of chromosomes in them; few other structures will be visible. They should aim to show the relative sizes and positions of the chromosomes and the cell accurately. Students should annotate the drawings to describe what is happening.


Technician Notes



Procedure continued

Part 2: MagnificationStudents use an eyepiece graticule to work out the magnification of the cells they have drawn.

20 30 40 50 60 70

1009080706050403020100

stage micrometerscale (each smalldivision usuallyequals 100 µm,so each largedivision is 1 mm)

eyepiecegraticule

scale

Figure B: Calibrating the stage micrometer using an eyepiece graticule

1. Students should calibrate the eyepiece graticule. They may either be told how to do this, or asked to research the process for themselves. This is the recommended procedure. Place a micrometer slide on the stage of the microscope. Using the low-power objective, focus on the micrometer scale. The smallest division of the micrometer scale is usually 100 μm. Move the slide and rotate the eyepiece to align the scales of the eyepiece graticule and the stage micrometer in the field of view. Count the number of divisions (eyepiece units or epu) on the eyepiece graticule that are equivalent to a known length on the micrometer slide and work out the length of each eyepiece unit. For example, if 100 μm is equivalent to 4 epu, then each epu is 100/4 = 25 μm at this magnification. Repeat this for the medium- and high-power objectives.

2. Students measure the length of the cells they have drawn. This is the image length.

3. They look at the original cell through the microscope and use the graticule to measure the length of the cell in micrometres.

4. They divide the image length by the actual length to calculate the magnification. They should add this value to their drawing.




Objective

To know how to carry out an investigation to determine the water potential of plant tissue

Safety ! Equipment

● Take care with glassware and cutting equipment. ● large potato

● thermometer

● 1 mol dm−3 sucrose solution

● distilled water

● boiling tubes and rack

● potato chip cutter

● 10 cm3 graduated pipette and pipette filler

● white tile

● digital balance

● scalpel

● ruler

● kitchen towels

● timer

● forceps

● marker pen

● access to a water bath

Background

There are two parts to this experiment: the first is to produce a dilution series and the second is to determine the water potential of plant tissue. Potatoes are used for this experiment, although it is possible to carry out a similar investigation using onions.

Procedure

Part 1: Preparing a dilution series1. Students use a marker pen to label six boiling tubes: 0.0, 0.2, 0.4, 0.6, 0.8 and 1.0 mol dm–3 sucrose. Depending

on time allocated, you may wish to prepare the dilutions in advance.

2. Students use the 1.0 mol dm–3 sucrose solution and water to make up 20 cm3 of sucrose solution in each concentration. They can then use the equation to work out the amount of sucrose solution. The water volume is the amount required to make up the total solution to 20 cm3. (Remind students they can also transfer this table to a spreadsheet to enable further analysis, for example to produce a graph.)

volume of sucrose solution required = required sucrose concentration

_________________________ starting sucrose concentration

× total volume of solution required

Concentration of sucrose solution (mol dm–3) 0.0 0.2 0.4 0.6 0.8 1.0

Volume of water (cm3) 20 16 12 8 4 0

Volume of 1.0 mol dm–3 sucrose solution (cm3) 0 4 8 12 16 20

Practical 3: Production of a dilution series of a solute to produce a calibration curve with which to identify the water potential of plant tissue

Technician Notes



Procedure continued

Part 2: Identifying the water potential of plant tissue1. Students place the boiling tubes containing the sucrose solutions in a water bath at 30 °C.

2. Six chips are cut from each student’s potato. If possible a chip cutter should be used to make sure all the chips have the same diameter. A ruler, scalpel and white tile can be used to cut all the chips to the same length. Make sure there is no skin on the chips.

3. Students blot the potato chips with kitchen towel until they no longer leave a wet patch on the towel.

4. The mass of each potato chip is then recorded in a results table.

5. After checking the solution in each boiling tube is at 30 °C using a thermometer, students place a potato chip in each boiling tube and start a stop clock.

6. After 30 minutes, the chips are removed and blotted dry, then reweighed and the new masses recorded.

7. The change in mass for each chip is then used to calculate the percentage change in mass.

Here is a typical conversion table for results.

Concentration of sucrose solution / mol dm−3 Water potential / kPa

1.00 −3500

0.80 −2580

0.60 −1800

0.40 −1120

0.20 −540

0.00 0

Practical 3: Production of a dilution series of a solute to produce a calibration curve with which to identify the water potential of plant tissue



Objectives

● To know how the effect of temperature on membranes can be determined

● To be able to recognise quantitative variables that should be controlled in an investigation

Safety ! Equipment

● Water baths at temperatures above 50 °C may scald. Take care when removing lids to allow steam to escape away from the face or body.

● Take care with sharp items such as the cork borer and knife.

● water baths pre-set at required temperatures

● thermometer ● distilled water ● beetroot ● cork borer ● ruler ● white tile ● knife

● 10 cm3 syringe ● pipette ● test tubes ● colorimeter ● cuvettes ● labels or pens for

labelling ● forceps ● crushed ice

Background

Students first carry out research to find out why beetroots appear red, what causes the red pigment to escape from cells and why this happens. Ensure they know to reference the sources they use correctly. They then carry out the investigation below.

Procedure

1. Students set eight water baths to a range of temperatures between 0 and 70 °C.

2. They then take eight test tubes and label each tube with the temperature of one of the water baths. A syringe is used to add 10 cm3 of distilled water to each test tube.

3. Each tube is placed in the water bath that is set to the corresponding temperature and left for 5 minutes.

4. Students check the temperature of each bath using a thermometer. The baths are unlikely to be at exactly the desired temperatures, so they should record the actual temperatures.

5. Students use cork borers to cut eight beetroot cylinders and then use a knife, ruler and white tile to trim the cylinders to the same length (1 cm is sufficient). The cylinders should be washed thoroughly with water until the water runs clear, and gently patted dry with a paper towel.

6. One beetroot cylinder is added to each test tube and left in the water bath for 15 minutes.

7. Students than shake each tube once. Working quickly, they should use forceps to carefully remove the cylinder from each tube. The cylinders are discarded, keeping the supernatant liquid (the clear liquid above the solid). It may be easier to decant the liquid into clean test tubes.

8. The colorimeter is set to a blue/green filter and percentage transmission, and zeroed using a blank cuvette filled with distilled water.

9. Liquid from each test tube in turn is transferred into a colorimeter cuvette, placed into the colorimeter and the percentage transmission reading is taken. Results are recorded in a suitable table.

white tile

beetroot

cork borerno. 4 or 5

ruler

Practical 4: Investigation into the effect of a named variable on the permeability of cell-surface membranes



Objective

To dissect, examine and draw the external and internal structure of the mammalian heart

Safety ! Equipment

● Wear a disposable apron to protect your clothing.

● Take care when using sharps to avoid cutting your hands.

● Wear gloves when handling hearts and wash your hands thoroughly after the practical. Dispose of used gloves immediately.

● disposable protective apron and disposable plastic/rubber gloves

● fresh sheep or pig heart

● wax dissection tray

● scissors

● mounted needle

● ruler or callipers with millimetre divisions

● paper towel

● blunt seeker

● disinfectant solution and plastic bags

● pins

● sticky labels

Note to teachers and studentsThis practical is based on a suggestion in the AQA Biology handbook. Students who do not wish to carry out a dissection of an animal on religious or moral grounds may study a plant organ or system instead.

Background

The heart is twisted so that the right ventricle (to your left as you dissect the heart) spirals behind and the left ventricle spirals to the front. Advise the students that to open the heart, it is better to make diagonal cuts following the coronary artery, since this artery follows the internal wall between the ventricles (the septum).

Procedure

atria

le� side of heart

coronaryartery

ventricles

Figure A: A diagram of a mammalian heart

1. Hearts should be placed on wax trays, with the left side of the heart to students’ right-hand side.

2. Students examine the coronary artery running across the surface of the heart and trace the artery up to the top of the heart and locate where it comes from the aorta. They may need to trim some fat away carefully.

3. Students now identify the two atria and the two ventricles and feel the difference in the thickness of the walls.

Practical 5: Dissection of animal or plant gas exchange or mass transport system or of organ within such a system

Technician Notes



Procedure continued

4. Using scissors, students pierce the left atrium wall, then cut down the heart in a straight line from the left atrium to the left ventricle. You may need to demonstrate this to your students.

5. The left atrium and the left ventricle are now opened to the apex of the heart.

6. Students identify the heart strings, or tendinous cords (chordae tendinae), and should observe their attachment via papillary muscles to the bicuspid valve and the ventricle wall.

7. Students trace where the aorta exits the left ventricle and identify the semilunar valve at the entrance to the aorta. The blunt seeker should be used to find openings and structures without cutting the tissue.

8. The thickness of the walls of the left atrium and the left ventricle should be measured in several places and the measurements recorded in a suitable table in the space provided in the lab book.

9. Students cut open the right side of the heart in the same way as the left.

10. The right side of the heart is now opened up, and the thickness of the atrium and ventricle walls measured. Measurements should be recorded in the space provided in the lab book.

11. Students identify the tricuspid valve, where the pulmonary artery exits the heart, and the semilunar valve.

12. Students draw diagrams of their dissections in the space provided in the lab book, annotating key features and adding a scale. They may also use pins and sticky labels to make flags to identify various parts of the heart, and could also take photographs of their diagrams to add to their notes.

13. Students annotate their drawings with the measurements they have recorded.

Practical 5: Dissection of animal or plant gas exchange or mass transport system or of organ within such a system



Objectives

● To observe the antibacterial properties of various substances

● To understand the aseptic techniques required in microbial tasks

Safety ! Equipment

● Follow aseptic techniques.

● Wash hands before and after handling the apparatus.

● Use a sterile plastic sheet to cover the surface of the clean bench prior to the experiment. Disinfect the bench after the experiment with 1% Virkon or equivalent. Leave the disinfectant on the bench for about 10 minutes.

● Wear a lab coat.

● McCartney bottle containing broth culture of Bacillus megaterium bacteria

● sterile plastic sheet on which to work

● 1% Virkon or equivalent ● prepared sterile agar

plates ● Bunsen burner ● agar slope or broth

cultures

● inoculating loop ● multodisc antibiotic

ring ● disinfectant ● paper towels ● marker pen ● sterile disposable

spreader ● autoclave tape ● sterile 1 cm3 pipette

and filler ● plastic forceps

Background

Students first carry out research into antimicrobials. Instruct the students to include material on disinfectants, antibiotics and antiseptics as part of their research. Ensure they know to reference the sources they use. They then carry out the investigation below.

Procedure

Part 1: Setting up plates 1. Remind students to thoroughly clean working areas, using disinfectant and wiping down with paper towels.

Plastic sheets should then be placed on the clean benches.

2. Students write their names, the date and the name of the bacteria on the underside of each agar plate.

3. The agar plates, the agar slope and the inoculating loop and placed next to a lit Bunsen burner.

4. Students should wash their hands before the next stage, and clear any spillages by mopping or placing over the spill a tissue soaked in 1% Virkon or equivalent.

5. Working close to the Bunsen burner, students should flame the neck of the McCartney bottle, then use the sterile pipette and filler to remove 0.3 cm3 of the bacterial culture. The neck of the McCartney bottle is then flamed again.

6. The agar plate should be opened as little as possible towards the flame and the contents of the pipette expelled onto the surface of the agar. The lid should be replaced on the plate and the used pipette placed in disinfectant, squeezing it first in the disinfectant to remove bacterial residue.

7. With the agar plate open as little as possible, the spreader is used to evenly distribute the bacteria across the surface of the agar. The lid should be replaced, and the spreader put in disinfectant.

8. Students use forceps to remove the multodisc (or mast ring) from its container and place it on the agar plate. Using the forceps, they should make sure the multodisc is in contact with the agar surface. The used forceps should be put in disinfectant.

9. Students seal the lid of the agar plate with two pieces of tape – being careful not to surround the lid with tape.

10. The plate is now incubated upside down for 24 hours at 25 °C.

11. Students should disinfect the bench and wash their hands thoroughly.

Practical 6: Use of aseptic techniques to investigate the effect of antimicrobial substances on microbial growth

Technician Notes



Practical 6: Use of aseptic techniques to investigate the effect of antimicrobial substances on microbial growth

Procedure continued

Part 2: Analysis of plates 1. Students must not open plates after incubation.

2. A ruler should be used to measure the diameter of zero growth – the zone of inhibition – around the antibacterial agent. This value can now be used to work out the area of the zone of inhibition.

BiologyTechnician Notes


AQA A Level

Objective

To separate and identify photosynthetic pigments from leaf chloroplasts

Safety ! Equipment

● Petroleum spirit is highly flammable, may cause severe lung damage if swallowed and is an irritant. Vapour may cause drowsiness or dizziness. Wear eye protection. Extinguish any naked flames close by. Do not inhale fumes, stopper containers and use in a well-ventilated area.

● Propanone is highly flammable and is an irritant to eyes and skin. Vapours may cause drowsiness and dizziness. Wear eye protection. Extinguish any naked flames close by. Do not inhale fumes. Use in small amounts in a well-ventilated area.

● pestle and mortar

● boiling tube and bung with pin

● boiling tube rack

● measuring cylinder

● spatula

● pencil and ruler

● fine tube for pigment loading

● leaves of different colours or from shade-tolerant and shade-intolerant plants

● chromatography paper ● propanone (pigment

extraction solvent) ● chromatography

solvent (1 : 9 mix of propanone : petroleum spirit)

Background

Chromatography is a separation process that relies on the differential distributions of a mixture between a mobile liquid phase (the chromatography solvent) and a stationary solid phase (in this case, paper). In this practical, students will investigate pigments either from leaves of different plants (i.e. leaves from shade-tolerant and shade-intolerant plants) or from leaves of different colours, depending on which leaves are available.

Procedure

level of solvent atend of experiment(solvent front)

distance travelled by solute (photosynthetic pigment)distance travelled by solvent

Rf value =

total distance travelledby solvent = x cm

baseline

distance travelled bypigment B = b cm

ax

a x

b

distance travelled bypigment A = a cm

ARf for =

bxBRf for =

Figure A: Separating pigments using chromatography

1. Students take a piece of chromatography paper of a suitable size to fit the full length of a boiling tube without touching the sides and draw a pencil line about 25 mm from the bottom edge.

Practical 7: Use of chromatography to investigate the pigments isolated from leaves of different plants e.g. leaves from shade-tolerant and shade-intolerant plants or leaves of different colours

Technician Notes


BiologyAQA A Level

Procedure continued

2. Wearing eye protection, students grind a few leaves in a mortar with a maximum of 10 cm3 of propanone to extract the pigments. The extract should be as concentrated as possible. Work should be done quickly to minimise evaporation.

3. Any pieces of plant material should be pushed to one side with a spatula. A fine pipette tip or capillary tube is then used to take up a small amount of extract. One small drop of this extract is then placed in the centre of the pencil line and allowed to dry before another drop is added on top. More drops are then added, allowing the pigment to dry each time, to build up a pigment spot that is as small as possible but dense enough that it contains sufficient pigment. Touching the chromatography paper with hands should be avoided, as fingerprints may interfere with the solvent movement. Students can begin Step 4 while the spots are drying.

4. Wearing eye protection, students carefully pour the chromatography solvent into a boiling tube to a depth of no more than 1 cm. Insert the bung.

5. When the chromatography paper is ready, it is suspended inside the boiling tube by pinning it to the underside of the bung or by trapping it in a split bung. The bottom of the paper should be dipped into the solvent, but the pigment spot must not be immersed in the solvent at any time.

6. The solvent front should rise up the paper, evenly separating different pigments. When it is close to the top of the tube (after around 10 minutes), the paper should be removed from the solvent and the position of the solvent front quickly marked using a pencil. The chromatogram is then allowed to dry in a well-ventilated area, preferably in the dark.

7. Steps 1–6 are repeated with a different leaf (i.e. a leaf with a different colour or tolerance of shade).

8. Once chromatograms are dry, students should examine them, and calculate the Rf value for each pigment using the procedure shown in Figure A.

Learning tip ● Remember that Rf values describe a ratio of the distance travelled by a pigment compared to the solvent front.

If a pigment travels half the distance of the solvent front, the Rf value will be 0.5. As they are ratios, Rf values are written without units.

Analysis of results1. Students record their results in the table provided, noting the colour of the pigment, the distance from the

baseline and the Rf value. They can then use their tables to identify pigment spots that have a similar Rf value.

Pigment Colour of spot Rf value

carotene yellow-orange 0.95

phaeophytin grey-yellow 0.83

xanthophyll yellow-brown 0.71

chlorophyll a blue-green 0.65

chlorophyll b light green 0.45

Table A: Appearance and Rf values for some common chloroplast pigments separated using propanone and petroleum spirit solvent

Practical 7: Use of chromatography to investigate the pigments isolated from leaves of different plants e.g. leaves from shade-tolerant and shade-intolerant plants or leaves of different colours



AQA A Level

Objective

To investigate the effect of ammonium hydroxide on dehydrogenase activity

Safety ! Equipment

● Very bright lights are needed for this practical. Some may emit UV light. Try not to look directly at the light.

● Take care with water next to electrical connections.

● Ammonium hydroxide is an irritant. Wear safety glasses.

● spinach leaves ● measuring cylinder ● filter funnel ● cold beakers ● test tubes ● test tube racks ● 1 cm3 and 5 cm3

syringes ● lamp ● marker pen ● timer

● blender ● muslin or similar

material for filtering ● DCPIP solution ● isolation medium ● 1.0 mol dm−3

ammonium hydroxide solution

● distilled water ● aluminium foil ● ice

Background

During the light-dependent reaction in photosystem I, NADP acts as an electron acceptor and is reduced (gains electrons). This reaction is catalysed by a dehydrogenase enzyme. DCPIP is a blue dye that can be used to monitor the rate of dehydrogenase activity. Any electrons released by the chlorophyll will change the colour of DCPIP from blue to colourless. The named factor being investigated is the action of ammonium hydroxide, which can be used to mimic the effect of weed killer.

Procedure

1. Students measure 50 cm3 of isolation medium into a cold beaker. It is likely that the isolation medium will be in a cold location (e.g. on ice or in a fridge).

2. Students take eight spinach leaves and tear them into small pieces. They place these pieces in the isolation medium in the cold beaker, ensuring only leaves are in the beaker, not stalks or midribs.

3. An already cooled large (250 cm3) beaker is half filled with ice. Students then label a smaller (100 cm3) beaker ‘chloroplast suspension’ and place it on top of the ice.

4. A piece of muslin is then wetted with isolation medium, and three layers placed over the top of a filter funnel. The filter funnel is then placed in the small beaker on ice.

5. Students now use a blender to blend the spinach and isolation medium.

6. The blended mixture is then returned to the original beaker.

7. The blended mixture is poured into the muslin in the filter funnel. All of the blended mixture needs to be filtered – the muslin may be squeezed if necessary to help with this filtering process.

8. Students label five test tubes 1, 2, 3, 4 and 5 and place these tubes in the ice in the large beaker.

9. A lamp is used to illuminate all the tubes.

Practical 8: Investigation into the effect of a named factor on the rate of dehydrogenase activity in extracts of chloroplasts

Technician Notes


BiologyAQA A Level

Procedure continued

10. Test tubes 1, 2 and 3 are set up based on the table below:

Test tube 1 Test tube 2 Test tube 3

DCPIP 5 cm3 5 cm3

water line above DCPIP 1 cm3 1 cm3 6 cm3

chloroplast suspension 1 cm3 6 cm3

isolation medium 1 cm3

aluminium foil cover completely

Test tubes 1 and 2 are control tubes and are left until the end of the investigation. Test tube 3 is a standard – it will show students the final colour for test tubes 4 and 5.

11. Test tube 4: Add 5 cm3 DCPIP, 1 cm3 water and 1 cm3 chloroplast suspension. The contents are mixed, and the time taken for the colour to change to green recorded. The colour change is complete when the colour matches that of test tube 3.

12. Step 11 is repeated four more times and the average time for the colour change worked out.

13. Test tube 5: Add 5 cm3 DCPIP, 1 cm3 ammonium hydroxide and 1 cm3 chloroplast suspension. The contents are mixed, and the time taken for the colour to change to green recorded. The colour change is complete when the colour matches that of test tube 3. If the colour has not changed completely after 5 minutes, students should record the colour at this point.

14. Step 13 is repeated four more times to work out the average time for the colour change. Solutions can be disposed of by diluting with lots of running water prior to placing down the sink.

Practical 8: Investigation into the effect of a named factor on the rate of dehydrogenase activity in extracts of chloroplasts



AQA A Level

Objectives

● To investigate the effect of different substrates on yeast respiration

● To conduct individual research into methods for measuring rate of respiration and the effect of different substrates on respiration

Safety ! Equipment

● If a kettle is used to heat water, take care with hot water.

● water bath at 40 °C or beaker with warm water

● thermometer

● four boiling tubes

● four Durham tubes

● pencil with rubber end

● four 2 cm3 yeast solutions, one for each substrate used

● 2 cm3 glucose solution

● 2 cm3 fructose solution

● 2 cm3 maltose solution

● 2 cm3 sucrose solution

● stop clock

● ruler

● chinagraph pencil

Background

Students are asked to combine the investigation below with individual research and to explain their findings in a report. Ensure students know to reference the sources they use.

Procedure

Procedure1. The apparatus is set up as shown in the diagram. Students fill the

beaker with warm water or use a water bath if you have already set one up.

2. Students place 2 cm3 yeast solution and 2 cm3 glucose solution in a Durham tube and cover with a boiling tube. They should use the rubber end of a pencil to hold the Durham tube firmly in place and then invert the boiling tube carefully, to ensure that none of the mixture escapes.

3. Students place the apparatus in the water bath and wait for the yeast to start respiring. When bubbles begin to appear in the Durham tube, they start the stop clock.

4. After 3 minutes a ruler should be used to measure the height of the gas produced in the Durham tube. The result should be recorded in the space provided in the lab book.

5. Students should now re-set the apparatus, this time using another substrate (fructose, maltose or sucrose) in place of the glucose, and repeat steps 3–4 for each substrate, recording the height of gas produced each time.

6. Students discuss among themselves and with their teacher how this apparatus could be modified to roughly measure the volume of gas produced. Once they have agreed a method, they write out the steps involved and then repeat the experiment using their modifications.

The rate of respiration with the most active substrate can then be calculated.

water bath

Durham tube with yeastand glucose solutions

pencil

Figure A: Use a pencil to push the fluid-filled smaller tube to the end of a boiling tube and invert

Practical 9: Investigation into the effect of a named variable on the rate of respiration of cultures of single-celled organisms



AQA A Level

Objectives

● To investigate the effect of an environmental variable on the movement of an animal

● To write a null hypothesis for an experiment

● To use the chi-squared statistical test in a practical setting

Safety ! Equipment

● Wash hands after handling woodlice.

● Do not eat the silica gel.

● a choice chamber with nylon mesh fabric

● silica gel ● black paper ● sticky tape ● humidity test strips

● paper towels ● water ● woodlice ● beaker ● forceps

Background

Typically, choice chambers are circular plastic boxes that are divided into areas which can be set up to have different conditions. In the lab, small invertebrates (usually woodlice) are placed in the chambers. After a period of time, the chambers are viewed and the location of the organisms is recorded. This provides information about the organisms’ ‘choices’ which can, hopefully, be explained by our expectations of the preferred environment for the organism being studied.

The two main sections of the choice chamber should be separated by a mesh. The woodlice will be introduced on top of the mesh through small holes in the upper container. The mesh will ensure that the woodlice are not in direct contact with any of the chemicals used to create the ‘mini’ environment.

Top chamber: includes holes to enable organisms to be easily introduced into the choice chamber

Lower chamber: normally where substances are added to a�ect the environment above

Mesh: o�en a fabric type material, allows organisms to experience a particular environment without direct contact with the substances causing the environmental change

Figure A: Setting up a choice chamber

Practical 10: Investigation into the effect of an environmental variable on the movement of an animal using either a choice chamber or a maze


Technician NotesPractical 10: Investigation into the effect of an environmental variable on the movement of an animal using either a choice chamber or a maze

Procedure

Part 1: Control1. The lower section of the choice chamber should be left empty.

2. Students put the mesh on top of the lower section.

3. The upper section is then placed on top of the lower section, with the mesh sandwiched between.

4. Ten woodlice are placed in the upper section through the holes.

5. Results should be recorded after a period of 5 minutes.

In this situation, it is expected that the left and right halves will have no effect on the distribution of woodlice: five woodlice will be expected in each half. However, this might not occur because of chance distribution. Using the chi-squared test (χ2) will determine the probability of the results occurring by chance. The experiment can proceed if the chi-squared test shows a greater than 5% probability of the results occurring by chance.

Part 2: Altering conditionsStudents can now investigate various conditions, including light, humidity or a combination of both.

To alter the light levels, cover sections with black paper. To alter the humidity, add damp paper towels or cotton wool to one section (making sure the mesh does not get wet) and add a small quantity of silica gel (a desiccant) to the other half, following the manufacturer’s instructions. Humidity levels will need to be tested before adding the woodlice to ensure there is a suitable difference between the sections.

BiologyAQA A Level



AQA A Level

Objectives

● To make a dilution series

● To produce a calibration curve

● To identify glucose in mock ‘urine’ samples

Safety ! Equipment

● Eye protection should be worn when using Benedict’s solution. Hands should be washed if the solution is spilt.

● Take care when handling the water bath and test tubes. There is a danger of scalding.

● 10% glucose solution ● Benedict’s solution ● graduated pipettes and pipette filler ● distilled water ● filter funnels ● filter paper ● colorimeter and cuvettes ● mock ‘urine’ samples ● test tubes ● test-tube rack ● water bath set

Procedure

Part 1: Glucose calibration curve1. Label six test tubes 1 to 6.

2. The test tubes are made up with the following mixtures, using the table below. Note that test tube 1 contains pure distilled water (plus Benedict’s solution) and test tube 6 contains the glucose solution supplied (plus Benedict’s solution).

Test tube 1 2 3 4 5 6

Dilution 0% glucose 0.5% glucose 1% glucose 2% glucose 5% glucose 10% glucose

Volume of glucose solution (cm3)

0 0.5 1 2 5 10

Volume of distilled water (cm3)

10 9.5 9 8 5 0

Benedict’s solution (cm3)

5 5 5 5 5 5

3. Students thoroughly mix the contents of the test tubes and put all the test tubes in a water bath set to 90 °C. The test tubes are left for 5 minutes, then removed from the water bath and allowed to cool in a test-tube rack.

4. Students use the contents of test tube 1 to calibrate the colorimeter to zero absorbance.

5. The remaining test tube solutions are now placed in cuvettes and their absorbance measured.

Part 2: Glucose concentration of urine samples1. Students make up a test tube for each urine sample, containing 10 cm3 ‘urine’ and 2 cm3 Benedict’s solution.

2. They then repeat steps 3–6 from Part 1 with each urine sample.

Practical 11: Production of a dilution series of a glucose solution and use of colorimetric techniques to produce a calibration curve with which to identify the concentration of glucose in an unknown ‘urine’ sample



AQA A Level

Objectives

● To investigate the effect of an abiotic factor on the distribution of a species ● To understand how to use transects to investigate the distribution of organisms

Safety ! Equipment

● Students must not throw quadrats. ● There is a low risk of infection from plants or soil, which

may be contaminated by animal faeces. Cover any cuts with a plaster and do not eat while working outdoors. Students should wash hands using soap after fieldwork.

● There is a possibility of allergic reactions to substances such as pollen, plant sap or insect stings. Ask students to inform you immediately if they feel unwell.

● Students should dress appropriately for wet and cold weather. They should wear sunscreen in summer months.

● If working near water, students must not enter water and must always work within sight of others.

● 0.25 m2 (0.5 m × 0.5 m) or 1 m2 quadrat with grid ● 20 m tape measure or rope marked at 1 m

intervals ● identification guide ● clipboard ● light meter or light sensor and data logger

Background

When investigating the difference between two environmentally distinct areas, random sampling within each area is appropriate. However, for changes in populations along an environmental gradient, a transect is a more appropriate method of sampling. Transect samples are taken systematically in a linear pattern. In this investigation students will use an interrupted belt transect to investigate the effect of one abiotic factor on the distribution of a single plant species. Light intensity is chosen as the abiotic factor in this example. An appropriate habitat to use might be a woodland area, a grassland area or a heath area.

Procedure

1. Students choose a site within their study area where there is an obvious gradient in the abiotic factor, for example, from a shaded area under a tree canopy or on the shaded side of a tall building to an area in full sunlight. They lay a 20 m tape or marked rope as a transect line with the 0 m mark in full shade and the 20 m mark in full light.

2. Students should walk along their transect, noting any change in species abundance. They choose which plant species to study, and decide whether to use percentage cover or counts of plants (density) as a measure of abundance. If counts are used, individual plants must be easy to distinguish and should not be too abundant to count easily.

3. Students lay the quadrat next to the 0 m mark of the tape. They record the abundance of their chosen species and measure the light intensity at ground level making sure not to shade the meter.

4. The quadrat is now moved 2 m along the tape and the light intensity measured again.5. Steps 3 and 4 are repeated until 10 measurements along the transect have been obtained. 6. Students repeat steps 3–5 with the tape positioned along two more transects running from a shaded area to

a lit area, to give repeat values at each distance. Alternatively, they can share class data.

Figure A: Transect sampling

Practical 12: Investigation into the effect of a named environmental factor on the distribution of a given species

Published by Pearson Education Limited, 80 Strand, London, WC2R 0RL.

www.pearsonschoolsandfecolleges.co.uk

Text © Pearson Education Limited 2018

Typeset and illustrated by Tech-Set Ltd Gateshead

Original illustrations © Pearson Education Limited

First published 2018

Copyright notice

All rights reserved. The material in this publication is copyright. Activity sheets may be freely photocopied for classroom use in the purchasing institution. However, this material is copyright and under no circumstances may copies be offered for sale. If you wish to use the material in any way other than that specified you must apply in writing to the publishers.

Acknowledgements

The publishers would like to thank John Kavanagh for his contributions to the text.

Note from the publisher

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