OCR A-level Biology Practicals

Practical 1 : Using a Light microscope to examine blood smears

Preparation:

1.) staining samples with methylene blue/ eosin

- staining to add contrast so it is easier to identify different parts of te cell

- for the electron microscope, the specimens are stained with lead

2.) mounting

- dry mount:

thinly sliced

use tweezers to pick up your specimen and put it in the middle of a clean slide

place a coverslip over the top of the specimen

- wet mount:

1.pipette a small drop of water onto the slide

2.use tweezers to place the specimen on top of the water drop

3.stand the coverslip upright on the slide next to the water droplet 4.and carefully tilt and lower the cover slide so it covers the 5.specimen to avoid air bubbles which may obstruct view.

6.stain the specimen by putting a drop of the stain next to the cover slip.

Microscope

1. clip the slide containing the specimen onto the stage

2. select lowest powered objective lens

3. use coarse adjustment knob to bring the stage up to just below objective lens

4. look down the eyepiece and the coarse adjustment knob to move the stage so the image is roughly in focus.

5. adjust the focus with the fine adjustment.

knob until a clear image is obtained

6. swap to greater magnification with higher-powered objective lens

REMEMBER

- MAGNIFICATION = IMAGE SIZE / OBJECT SIZE

- stage micrometer is the bigger 'ruler'

- eye piece graticule is the smaller 'ruler'

Practical 2 : Dissection of a mammalian heart

equipment: pig's heart, dissecting tray, scalpel, spron and lab gloves

external examination:

- try to identify four main vezzels

-arteries are thick and rubbery

- veins are thinner

- identify the right and left atria

internal examination

- cut along the lines

- measure and record the thickness of the ventricle walls note any differences ventricular walls

- cut open the atria and look inside them too.Note whether atria walls are thicker or thinner than the ventricular walls

- find AV valves and semi-lunar valves.

- Draw sketch to show the valves

Practical 3 : Dissection of a stem

1. use a scalpel to cut a cross-section of the stem (transverse or longitudinal) as thinly as possible

2. use tweezers to gently place the cut sections in water

3. transfer each section to a stain toluidine blue o, stains the ligning in xylem walls blue-green, identify xylem structure

4.rinse off the sections(so the pigment does not obstruct the speciment0 and mount onto a slide to view under the microscope

Practical 4: Calculation of species diversity /Practical 5: Measurement of the distribution and abundance of plants in a habitat

Comparing the abundance of plant species in two different fields

1. choose and area to sample- within the habitat being stidues

2. count the number of individual of each species (specirs eveness)

- plants: quadrats, transect line

- flying insects: sweep net

-ground insects: pitfall trap

- aquatic animals: net

3. use the results to estimate the total number of individuals or the total number of different species in the habitat being studied.

4. when sampling different habitats and comparing them, ALWAYS USE THE SAME SAMPLING TECHNIQUE.

3

*avoid bias: random sampling ( random generation of coordinates coordinates on a grid made by two transect lines)

* Non-random sampling

1. systematic: samples are taken at fixed intervals (often along a transect line)

2. Opportunistic: samples are chosen by the investigator, simple to carry out, biased data.

3. Stratifies- different areas in a habitat are identified and samples separately in proportion to their part of the habitat as a whole.

genetic diversity is measured by proportion of polymorphic gene loci = number of polymorphic gene loci / total number of loci

species richness: the number of different species living in a particular area

species evenness- a comparison of the numbers of individuals of each species living in a community

Investigating the distribution of plant species along an abiotic gradient

Practical 6: The effect of substrate/enzyme concentration on enzyme controlled reaction *catalase

For investigating different enzyme and substarte concentrations.

1. set up boiling tubes containing same volume and concentration of substrate (or enzyme)

2. set up boiling tubes containing different concentrations of enzymes (or substrate)

3. add equal volumes of buffer solution

4. set up apparatus to measure the gas given off for colour change

5. time how long it takes for changes to happen

* for temperature investigation use water bath

* for pH investigation pre-measure pH and use different pH for each test (don

*keep pH and temperature constant

Practical 7: Investigating the effect of temperature on amylase activity

1. add equal volumes of iodine drops (amylase breaks down starch) into dropping tiles

equal volumes of starch solution and amylase

2. same volume and concentration of amylase and starch solutions incubated in a water bath at set intervals on temperature for the same amount of time until the desired temperature is reached

3. add amylase enzyme to starch and iodine

4.time how long it take for when the iodine solution in the spotting tile remains orange-brown (no starch is being broken down).

variable that need to be controlled

- pH

- temperature

-enzyme concentration

- substrate concentration

When enzyme from natural source how to control experiment better

-same plant

-same part of plant

-same mass of tissue

-same size

*Practical 8: The effect of temperature on membrane permeability (beetroot experiment)

1. cut equal size pieces of beetroot and rinse them to remove any pigment released during cutting.

2. Place the five pieces in five different test tubes, each with the same volume of water

3. Place each test tube in a water bath set at different temperatures for the same length of time

4 remove the pieces of beetroot from the test tubes, leaving just the coloured liquid

5. take the small volume of each coloured liquid and put it into covet for the colorimeter

*make sure to calibrate the colorimeter with a blank first

6. measure the absorbance rate of each liquid

7. the more absorbance (less light passing through) the more permeable the membrane was

* more transmission = less permeable the membrane

variables to control:

-temperature

-ethanol concentration

-No excess dye on cut sections of beetroot (rinse off)

-swirl liquid before putting in the cuvette for the

-time

-beetroot (age, same plant, same part)

-SA and/or mass of beetroot used.

** remember

judging colors by eye is not as good as using colorimeter because the results are judged subjectively or by using a colorimeter

Practical 9: Determining glucose concentration

Method 1: Test strips and then compare to a chart

Practical 10: Using a potometer (Factors affecting transpiration rates)

1. cut a shoot underwater to prevent air from entering the xylem (interferes with the column of wtaer travelling up the xylem) cut a slant to increase the surface area available for water uptake.

2. Assemble the potometer in the water and insert the shoot underwater, so no air can enter the water.

3. remove the apparatus from the water but keep the end of the capillary tube submerged in a beaker of water.

4. check the apparatus is watertight (vaselline etc)

5 dry the leaves to allow time for shoots to acclimatize

6. remove the end of the capillary tube from the beaker of water until one air bubble had formed, put the capillary tube back into the water.

7. record the starting position of the air bubble along the ruler.

8. start the stopwatch and record the distance moved by the bubble per unit time

9. rate of air bubble movement is the rate of transpiration

10. all other conditions must be kept constant. only one change should be variable.

independent variables:

- wind speed

- use a fan

-humidity (plastic bag over the plant)

-light intensity ( lamp distance away from plant)

-temperature

variables to control

water uptake

no all water is ivolved in transpiration

some may be used to maintain tugidity or some use in photosynthesis

Practical 11: The effect of antibiotics on bacterial growth

1. dip discs in antibiotics

2. place on bacterial culture

3. measure inhibition zone

Practical 12: Dilution plating to determine the density of microbes in liquid culture. **biotechnology and bacterial culture. (Factors affecting the growth of microorganisms)

1. supplies a sample of bacteria broth.

2 make dilutions if necessary

3. using a sterile pipette add a set volume of sample onto agar plate

4. spread the brith across entire surface of the agar using a sterile spreader

5. put the lid on the agar plate and tape it shut

repeat steps 3-5 for six plates

control : uncultured agar plate

leave all plates for the same amount of time

6. count the number of colonies that have formed on each pkate

7. work out the mean number f colonies formed under each condition

Practical 13: Water potential of potato (The effect of solutions of different water potentials on plant/animal cells)

1. prepare sucrose solutions at different concentrations

2. use a cork borer or chip maker to cut potatoes into same-sized pieces

3. dry the potato chips with paper towel (same method for a potato cylinders)

4. measure the mass of each potato chip and record it

5. place one potato chip in each solution

6 leave the chips in solution for the same amount of time

7. remove the chips and pat gently with a paper towel

8. re-weigh each group again and record results

9. calculate the percentage change in mass for each potato chip

10. plot results on graph

Independent variable

-Water potential of the surrounding liquid

- type of cell

variables to control

-time

-mass

-size

- surface area of tissue

- same plant/ tissue (part of plant)

- potato/ visking tubing is dry

** the points at which the graph crosses the x-axis is the isotonic point

Practical 14: Osmosis in an artificial cell

like a potato but with different water potential in artificial cell and in surrounding solutions and no need to dry the

Practical 15: Rate of diffusion through a membrane (Factors aff ecting diffusion in model cells)

1. make up agar jelly with phenolpthalein and dilute sodium hydroxide

2. fill a beaker with some dilute hydrochloric acid

3. cut out cubes of different dimensions

4. time how long it take for cubes to go colourless

5. calculate the rate at which it took for the colour to disappear

OR

cut cubes the same size but submerge in different concentrations of HCl

variables to control

- temperature

-size and shape of agar blocks

Practical 16: Qualitative testing for proteins

1. add a few drop sodium hydroxide solution

2. add equal volume of biuret solution

if protein is present the solution will turn from blue to purple

Practical 17: qualitative testing for lipids )Emulsion test

1. Add equal volume ethanol

2. shake the solution

3. add solution into distille water

if lipid present their will e a milky emulsion

Practical 18: Qualitative testing for sugars (reducing and non-reducing)

reducing sugars (all monosaccharide and dissacharides except sucrose)

1. Add the same volume Benedict's reagent to s ample and heat it in a boiling water bath

2.if test is positive then coloured precipitate will form on spectrum blue to red

non-reducing sugars

1. add dilute hydrochloric acid

2, heat in a boiliing water bath

3.neutralise with sodium hydrogencarbonate

4. carry out benedicts test

5 if results are positive then you have a reducing sugar

Practical 19: Test for starch

1. add iodine

2. if starch is present it will turn from browny-orange to blue-black colour

Practical 20: Investigation into effect of exercise on pulse rate

Investigation into phototropism

1. Take nine wheat shoots. The shoots should be planted in individual pots in the same type of soil. The shoot should be roughly equal in height.

2. Cover the tips of the three shots with foil cap. Leave three shoots without foil. Wrap the base of the ifnal three shoots with foil, leaving only the tip expose

3. set up the shoots in front of the light source and leave them for two days. The shoots should all be the same distance from the lights source and experience the same intensity of light. All other variables including temperature and exposure to moisture, should be controlled.

4.By the end of the experiment you should be able to observe how the plans have changed in response to the light. The shoots with exposed tips should have grown towards the light source (positive tropism). Covering the tip with a foil cap prevents growth towards the light, it's the tip that's most sensitive to light and because it's covered the shoot should have continued to grow straight up. Covering the base of the shoot with foil should still allow the tip to grow towards the light.

5. Recording the amount of growth (in mm) , as well as the direction of growth, will give you quatitative results.

Investigations into Geotropism too

1. Line three Petri dishes with moist cotton wool.

You should use the same volume of water and the same amount of cotton wool in each dish

2. Space 10 cress seeds on the surface of the cotton wool in each dish, then push each one into the wool.

3. Tape a lid onto each dish and wrap each one in foil (this will prevent any light reaching the seeds and affecting your results).

4. Choose somewhere you can leave this dishes where the temperature is likely to be warmish and pretty constant e.g. cupboard.

5. Prop one dish. upright , at a 90 angle - label it and mark which way is 'up' (or down) . Place another dish on a slope at a 45 angle. Place the third dish on a flat, horizontal surface. You need to label the dishes carefully so you know which way up each one was when you come to unwrap the dishes at the end of the experiment.

6.Leave the seeds for 4 days. Then take a look at their shoot and root growth.

7. You should find that whatever the angle the dishes were placed at, the shoots have all grown away from gravity (negative geotropism) and the roots have grown towards gravity (positive geotropism).

8. To get quantitative results, measure the amount of growth of the shoots and root and the angle of growth.

Practical 21: Investigation into plant hormones (Auxin)

1. Plant 30 plants that are similar age, height and weight in pots

2. Count and record the number of side shoots growing from the main stem of each plant.

3. For 10 plants, remove the tip of the shoot and apply a paste without auxins to the top of the stem.

4. For another 10 plants, remove the tip of the shoot and apply a paste with auxins to the top of the stem

5. Leave the final 10 plants as they are - these are your controls. They are needed so you know that any effects observed is caused by the application of plant hormone auxin.

6. Let each group now grow for six days. You need to keep all the plants in the same conditions- the same, light intensity, water, temperature, etc. This ensures any variable that may affect your results are controlled, which makes your experiment more valid.

7.After six days, count the number of sides shoots growing from the main stem of each of your plants.

8. The results in the table show that removing the tips of shoots caused extra side shoots to grow, but removing tips and applying auxins prevented extra side shoots from growing.

9. the results suggest auxins inhibit the growth of side shoots- suggesting that auxins are involved in apical dominance.

Practical 21: Investigation into plant hormones (Gibberlins)

1. Plant 40 plants that are similar age, height and weight in pots

2. Leave 20 plants as they are to grow, watering them all in the same way and keeping them all in the same conditions - these are your controls.

3. Leave the other 20 plants to grow in the same conditions except a dilute solution of gibberlin.

4. Let the plants grow for about 28 days and measure the lengths of all the stems once each week.

5. You light get results a bit like these:

6. The results in the table show that stems grow more when watered with a dilute solution of gibberlin

7. The results suggest gibberlin stimulate stem elongation.

8.May have to calculate the rate of growth such as cm/day

Investigating the rate of respiration of yeast under aerobic and anaerobic conditions

AEROBIC

1. Put a known concentration of substrate solution (e.g. glucose in the test tube

2. Add a known volume of buffer solution to keep the pH constant (remember to choose the optimum pH for the yeast you are testing usually around pH 4.6)

3.Place the test tube in a water bath set to 25 degrees celsius. This ensures the temperature of the substrate to stabilise. Leave for 10 minutes to allow temperature of substrate to stabilise.

4.Add a known mass of dried yeast to test tube and stir for two minutes.

5. After the yeast has dissolved into the solution, put a bung with a tube attached to the gas synringe in the top of the test tube. The gas syringe should be set to zero

6. Start the stopwatch as soon as the bung has been put into the test tube.

7. As the yeast respire the carbon dioxide formed from the respiration will travel up the gas synringe used to measure the volume of CO 2 produced.

8. At regular time intervals record the volume of gas produced (present in the gas syringe). Do this is for a set amount of time.

9. A control experiment should also be set up where no yeast is present. expected results in the control would be that no carbon dioxide will be produced.

10. Repeat the experiment three times. Use your data to calculate a mean of CO2 production

ANAEROBIC

1. Set up the test tube with gas syringe apparatus

2. After the yeast is dissolved into the substrate solution, trickle some liquid paraffin down the side of the test tube so that it settles on and completely cover the surface of the solution. This will stop oxygen getting in forcing the yeast to respire anaerobically.

3. Put a bung of the tube attached to a gas syringe in the top of the test tube. The gas syringe should be set to zero.

4. perform steps 6-10 from the aerobic respiration experiment above.

Practical 22: Investigation into the respiration rate of yeast (Factors affecting the rate of respiration) ***respirometer

1. Set up the apparatus like on the diagram

2. Each tube contains potassium hydroxide solution (or soda lime) to absorb the carbon dioxide produced.

3.The control tube is set up in the exactly the same as the test tube, but without the woodlice, this is to make sure that any results observed are only due to the woodlice respiring. The control will usually contain some beads that are the same mass as the woodlice.

4. coloured fluid is then added to the manometer by dipping the end of the capillary tube into a beaker of fluid. Capillary action will make the fluid move into the tube. The syringe is then used to set the fluid to a know level.

5. The apparatus is left for a set period of time

6.During that time there'll be a devrease in the volume of air in the test tube due to the oxygen consumption of the woodlice ( all the Co2 produced is absorbed by the potassium hydroxide)

7. The decrease in the volume of air will reduce the pressure in the tube and cause the coloured liquid in the manometer t move towards the test tube

8. The distance moved by the liquid in a given time is measured. This value can then be used to calculate the volume of oxygen taken in by the woodlice per minute.

* remember you need to know the diameter of the capillary to do this.

9. Any variables that could affect results are controlled e.g. temperature, volume of potassium hydroxide solution in the test tube.

10. To produce more precise results, the experiment is repeated and mean volume of O2 is calculated.

Practical 23: Investigation into the rate of oxygen production in pondweed (photosynthesis). light intensity (Factors affecting the rate of photosynthesis)

1. A test tube containing pond weed and water is connected to a capillary tube full of water

2. the capillary tube is connected to a syringe

3. a source of white light s place at a specific distance from the pondweed.

4. The pondweed left to photosynthesise for a set amount of time. As it photosynthesises, the oxygen released will collect in the capillary tube.

5. At the end of the experiment, the syringe is used to draw the gas bubble in the tube up alongside the ruler and the length of the gas bubble is measured. This is proportional to the volume of O2 produced.

6. Any variable that could affect the results should be controlled. e.g. the temperature the time the weed is left to photosynthesise.

7. The experiment is repeated and the average length of gas bubbles calculated, to make the results more precise.

8. The whole experiment is then repeated with the light source placed at different distances from the pondweed.

9. to make the control use a test tube with no pondweed

Paper/Thin-Layer Chromatography

1. Grind up several leaves with anhydrous sodium sulfate and propanone.

2. Transfer the liquid to test tube, add some petroleum ether and gently shake the tube. Two distinct layers form in the liquid, the top layer is the pigments mixed in with the petroleum ether.

3. Transfer some of the liquid from the top layer into a second test tube with some anhydrous sodium sulfate.

4. Draw a horizontal pencil line near the bottom of a chromatography plate (silica plate). Build up a single concentrated spot of the liquid on the line by applying several drops and ensuring each one is dry before the next one is added. This is the point of oriigin.

5.Once the point of origin is completely dry, put the plate into a glass beaker with some prepared solvent ( eg. a mixture of propanone, cyclohexane, and petroleum ether. Lower is just enough so that the point of origin is slightly above the solvent. Put a lid on the beaker and leave the plate to develop. As the solvent spreads up the plate, the different pigments move with it, but at different rates - so they separate.

6.When the solvent has nearly reached the top, take the plate out and mark the solvent front (the furthest point the solvent has reached) with a pencil before it evaporates and leave the plate to dry in a well-ventilated place.

7. There should be several new coloured spots on the chromatography plate between the point of origin and the solvent front. These are the separated pigments. You can calculate the rF values and look them up in a data base to identify the pigments.

Computer modelling and data logging

COMPUTER MODELLING

advantages:

- Allows manipulation and measurements to be taken that couldn't be done with actual molecules.

-Allows the potential to simulate interactions between a range of different molecules at the atomic level.

-At a broader level computer modelling allows whole systems (e.g. metabolic pathways, ecosystems, climate) to be investigated where experiments are not possible

-Models can make predictions about the impact of changes in the system that may not be testable experimentally.

disadvatanges:

-Can require a large amount of data processing (high cost and too slow).

- Accurate and valid data needs to be collected to build a valid model

- Not all factors that affect a system can be included in a model which will affect validity.

-Predictions that models make are only valid if the assumptions of the model are true and if no unforeseen changes occur in the future.

DATA LOGGING

advantages:

-Can be done remotely and continuously.

-Data can be easily stored for future analysis or transmitted elsewhere foranalysis.

-Often the equipment is more precise than alternative ways of measuring.

disadvantages:

-Equipment can be expensive

-If data is being collected remotely, and the data logger retrieved at the end of data collection for downloading all the data could be lost if the data logger is lost or damaged.

DNA extraction

grind a sample in pestle and mortar

mix detergent with the sample - breaks down the cell membrane

add salt - break hydrogen bonds

add protease enzymes- brekadown histone proteins

add ethanol cause dna to precipitate out of the solution

DNA will be seen as white strands forming bewteen the layer of sample and layer of alcohol