Biology 9700/38 — May/June 2025

Background Concept

A serial dilution is a standard laboratory technique used to prepare a range of solutions of decreasing concentration. Each successive solution is made by diluting the previous one by a fixed factor. When the dilution factor is 2 (i.e. halving the concentration each time), the volumes transferred and added must be equal: transferring $V\ \text{cm}^3$ of the previous solution and adding $V\ \text{cm}^3$ of diluent (distilled water W) gives a total of $2V\ \text{cm}^3$ at half the original concentration. This is the simplest and most common form of serial dilution, and it gives a geometric series of concentrations (1, 1/2, 1/4, 1/8, 1/16, ...).

Understanding the Question

Fig. 1.2 shows the start of a serial dilution: the first beaker contains $20\ \text{cm}^3$ of 1.0% copper sulfate solution C, and the candidate is asked to extend the dilution through four more beakers so that five concentrations (1.0% plus four new ones) are available. Each beaker should contain a labelled arrow showing the volume of copper sulfate solution transferred, a labelled arrow showing the volume of distilled water W added, and a label showing the concentration under the beaker. The end result must give $10\ \text{cm}^3$ of each concentration available for the experiment.

Approach

To halve the concentration at each step while still leaving $10\ \text{cm}^3$ of usable solution in the previous beaker, transfer $10\ \text{cm}^3$ from the previous beaker and add $10\ \text{cm}^3$ of distilled water W. The new beaker will contain $20\ \text{cm}^3$ at half the concentration. The candidate then uses $10\ \text{cm}^3$ of this in the experiment and transfers $10\ \text{cm}^3$ to the next beaker. The concentrations halve each time: 1.0% → 0.5% → 0.25% → 0.125% → 0.0625%.

Step-by-Step Reasoning

1. Beaker 1 (already given): $20\ \text{cm}^3$ of 1.0% CuSO$_4$ + $0\ \text{cm}^3$ water = $20\ \text{cm}^3$ at 1.0%. Of this, $10\ \text{cm}^3$ is used in the experiment and $10\ \text{cm}^3$ is transferred to beaker 2.
2. Beaker 2: $10\ \text{cm}^3$ of 1.0% CuSO$_4$ + $10\ \text{cm}^3$ water = $20\ \text{cm}^3$ at $\frac{10 \times 1.0\%}{20} = 0.5\%$. Of this, $10\ \text{cm}^3$ is used in the experiment and $10\ \text{cm}^3$ is transferred to beaker 3.
3. Beaker 3: $10\ \text{cm}^3$ of 0.5% CuSO$_4$ + $10\ \text{cm}^3$ water = $20\ \text{cm}^3$ at 0.25%. Of this, $10\ \text{cm}^3$ is used and $10\ \text{cm}^3$ is transferred to beaker 4.
4. Beaker 4: $10\ \text{cm}^3$ of 0.25% CuSO$_4$ + $10\ \text{cm}^3$ water = $20\ \text{cm}^3$ at 0.125%. Of this, $10\ \text{cm}^3$ is used and $10\ \text{cm}^3$ is transferred to beaker 5.
5. Beaker 5: $10\ \text{cm}^3$ of 0.125% CuSO$_4$ + $10\ \text{cm}^3$ water = $20\ \text{cm}^3$ at 0.0625%. All $10\ \text{cm}^3$ is used in the experiment (no further beaker).

The final answer for Fig. 1.2 is four beakers (2 to 5), each showing a curved arrow from the previous beaker labelled "$10\ \text{cm}^3$ of copper sulfate solution", a vertical arrow into the beaker labelled "$10\ \text{cm}^3$ of distilled water W", and a concentration label underneath (0.5%, 0.25%, 0.125%, 0.0625%).

Key Takeaways

• A serial dilution halves the concentration when equal volumes are transferred and added.
• Each beaker in this series holds $20\ \text{cm}^3$, providing $10\ \text{cm}^3$ for use and $10\ \text{cm}^3$ to pass on (except the last, where all $10\ \text{cm}^3$ is used).
• The five concentrations are 1.0%, 0.5%, 0.25%, 0.125% and 0.0625%.

Common Mistakes

• Using unequal volumes (e.g. transferring $5\ \text{cm}^3$ and adding $15\ \text{cm}^3$ would dilute by a factor of 4, not 2).
• Labelling the wrong concentration under a beaker, or skipping a value in the sequence.
• Forgetting to include the volume labels on the arrows.
• Putting the wrong total volume in the beakers (each must end with $20\ \text{cm}^3$ for the procedure to work).

Things to Be Careful About

• The candidate must show $10\ \text{cm}^3$ transferred (not $20\ \text{cm}^3$); if all $20\ \text{cm}^3$ were transferred, the previous beaker would be empty.
• The candidate must add $10\ \text{cm}^3$ of water W to each of beakers 2 to 5 (not $0\ \text{cm}^3$, which is correct only for beaker 1).
• The concentration labels must appear in the correct order along the diagonal: 0.5%, 0.25%, 0.125%, 0.0625%.

Background Concept

A results table should have a clear heading for each column that includes both the quantity being measured and its unit. The independent variable (the one deliberately varied by the experimenter) is placed in the left column, and the dependent variable (the one measured) in the right column. If the same procedure is repeated, replicates can be added as additional columns. Here the candidate records a single set of readings (one per concentration), so two columns are sufficient.

Understanding the Question

The candidate has carried out the procedure in step 1 to step 14 and now needs to record, in an appropriate table, the number of drops of potassium manganate(VII) P needed to reach the end-point for each of the five concentrations of copper sulfate prepared in step 1. The independent variable is the concentration of copper sulfate; the dependent variable is the number of drops of P. The mark scheme requires:

• a heading for the independent variable to the left of the dependent variable
• a heading for the dependent variable (number of drops of P)
• a value for each of the five concentrations
• whole drops only
• the expected trend (the higher the concentration of CuSO$_4$, the more drops of P)

Approach

Construct a two-column table with the concentration of copper sulfate (in %) on the left and the number of drops of P on the right. Record one whole number per row, in ascending or descending order of CuSO$_4$ concentration. The expected trend reflects the inhibition: as CuSO$_4$ increases, less catalase is active, less H$_2$O$_2$ is broken down in the 5 minutes, and so more P is needed to react with the remaining H$_2$O$_2$.

Step-by-Step Reasoning

1. Identify the independent variable: concentration of copper sulfate (%). Place in the left column with the unit.
2. Identify the dependent variable: number of drops of P. Place in the right column.
3. List the five concentrations: 0.0625%, 0.125%, 0.25%, 0.5%, 1.0% (or in reverse order).
4. Record the number of drops of P needed to reach the end-point for each concentration. Each value is a whole number (or 'more than 30' if the end-point is not reached within 30 drops).
5. Check the trend: as CuSO$_4$ concentration rises, the number of drops of P should rise. If the trend is reversed, suspect a procedural error (e.g. the CuSO$_4$ may have been added in the wrong order, or the same pipette may have been used between concentrations without rinsing).

Key Takeaways

• A table of results should have a clear heading for each column including the unit.
• The independent variable is on the left; the dependent variable is on the right.
• In an inhibition experiment, more inhibitor means more substrate remains, so the dependent variable (related to substrate concentration) should rise with the inhibitor.

Common Mistakes

• Omitting the unit from the column heading (e.g. just 'concentration of copper sulfate' without the %).
• Putting the dependent variable (drops of P) to the left of the independent variable (concentration of CuSO$_4$).
• Recording non-integer values (e.g. 14.5 drops) when the procedure specifies whole drops.
• Recording a trend that decreases with concentration (which would suggest the inhibitor is acting as an activator — not what the data show).

Things to Be Careful About

• Whole drops only — the procedure involves counting each drop, so fractional drops cannot be recorded.
• 'More than 30' is an acceptable entry if the end-point is not reached; the mark scheme accepts this.
• The order of rows can be ascending or descending concentration; either is acceptable as long as the trend is clear.

Background Concept

Catalase is an enzyme in yeast cells that breaks down hydrogen peroxide (H$_2$O$_2$) into water and oxygen. Copper sulfate is a known inhibitor of catalase: at higher concentrations, the active site of the enzyme is more likely to be occupied by copper ions, reducing the rate at which catalase breaks down H$_2$O$_2$. After a fixed time (5 minutes), if more H$_2$O$_2$ remains, it means less breakdown has occurred, i.e. the enzyme has been more inhibited.

Understanding the Question

The candidate has measured the number of drops of P needed to reach the end-point for each CuSO$_4$ concentration. Because more drops of P are needed when more H$_2$O$_2$ is present, the number of drops is a direct indicator of the H$_2$O$_2$ concentration remaining after 5 minutes. The question asks the candidate to describe the relationship between CuSO$_4$ concentration and H$_2$O$_2$ remaining.

Approach

State the trend in plain words: as CuSO$_4$ increases, H$_2$O$_2$ remaining also increases. The mark scheme expects a single sentence linking the two variables in the correct direction.

Step-by-Step Reasoning

1. Look at the table: as CuSO$_4$ increases from 0.0625% to 1.0%, the number of drops of P increases (e.g. 7 → 25 in the representative table).
2. More drops of P means more H$_2$O$_2$ is present, because each drop reacts with a fixed amount of H$_2$O$_2$ until the end-point is reached.
3. Therefore, more H$_2$O$_2$ remains at higher CuSO$_4$ concentrations. This reflects greater inhibition of catalase.

Key Takeaways

• A positive correlation between CuSO$_4$ concentration and the number of drops of P means a positive correlation between CuSO$_4$ concentration and the H$_2$O$_2$ remaining.
• The biology: copper sulfate inhibits catalase, so the enzyme breaks down less H$_2$O$_2$ in 5 minutes when more inhibitor is present.

Common Mistakes

• Stating the wrong direction (e.g. 'the higher the CuSO$_4$, the lower the H$_2$O$_2$ remaining') — this is the opposite of what the experiment shows.
• Being vague ('CuSO$_4$ affects H$_2$O$_2$') without specifying the direction.
• Confusing the cause and effect (e.g. saying 'more H$_2$O$_2$ causes more CuSO$_4$' instead of the other way around).

Things to Be Careful About

• The question asks for a description, not an explanation. A single sentence stating the trend is sufficient.
• The exact phrasing matters: the mark scheme looks for 'the higher the concentration of copper sulfate, the higher the concentration of hydrogen peroxide'. A paraphrase that captures the same idea is acceptable.

Background Concept

In a drop-counting titration, the end-point is reached when enough drops have been added to react with all of the substance being measured. The accuracy of the method depends on each drop being the same size and the end-point being detected reliably. Any variability in drop size or difficulty in detecting the colour change introduces error into the count.

Understanding the Question

The candidate is asked to describe ONE source of error in steps 10 to 12 of the procedure, which involve releasing drops of P from a syringe one at a time into the test-tube and observing when the pink colour persists for at least 5 seconds. The mark scheme accepts three specific answers.

Approach

Think about what could go wrong when adding drops one at a time and judging the colour. The most common issues are: (1) the size of drops from a syringe is not perfectly uniform, (2) it is hard to release exactly one drop at a time, and (3) the end-point colour (pink persisting for 5 seconds) can be hard to judge.

Step-by-Step Reasoning

1. Drop size: a syringe does not deliver perfectly uniform drops; small variations in the diameter of the drop or in the force applied to the plunger can change the volume. Different drops therefore contain slightly different amounts of P, so the count is not perfectly proportional to the amount of H$_2$O$_2$ present.
2. Drop-by-drop control: releasing exactly one drop is difficult; sometimes two drops come out together, or the drop hangs from the tip and falls later, disrupting the count.
3. End-point detection: the pink colour may be hard to see against the background or in the small volume of the test-tube, and judging when the colour 'stays for at least 5 seconds' introduces a subjective time element.

Key Takeaways

• Sources of error must be specific, not vague. 'Human error' or 'not accurate' is too general and is not credited.
• Drop-counting methods have inherent limitations: drop size varies, single-drop release is difficult, and the end-point is subjective.

Common Mistakes

• Giving a vague answer like 'human error' or 'the experiment might not be accurate'.
• Naming an error that is not in the procedure (e.g. the temperature changing, when the procedure does not involve heating).
• Stating a source of error without linking it to the drop-counting step.

Things to Be Careful About

• Only ONE source of error is required; the mark is for a single, specific, plausible error.
• The error must be in steps 10 to 12 (the drop-counting steps), not in step 1 to 7 (the dilution and reaction steps).

Background Concept

The same drop-counting procedure used for the standard CuSO$_4$ concentrations in (a)(ii) is now applied to the river water sample R, which contains an unknown concentration of CuSO$_4$. The procedure is identical: $1\ \text{cm}^3$ of R is mixed with $1\ \text{cm}^3$ of yeast Y, then $1\ \text{cm}^3$ of H$_2$O$_2$ H, and after 5 minutes the reaction is stopped with sulfuric acid A; the number of drops of P needed to reach the end-point is recorded.

Understanding the Question

The candidate is asked to record the number of drops of P needed to reach the end-point for the river water sample R, using the same procedure as in steps 5 to 12. The mark scheme accepts any whole number (or 'more than 30' if the end-point is not reached).

Approach

Carry out the procedure with R and count the drops. Record a whole number. The mark scheme requires only that the value be a whole number, not a specific value.

Step-by-Step Reasoning

1. The candidate has labelled a test-tube R and added $1\ \text{cm}^3$ of R to it.
2. The same procedure as in steps 5 to 12 is then followed: $1\ \text{cm}^3$ of Y, wait 2 minutes, $1\ \text{cm}^3$ of H, wait 5 minutes, $1\ \text{cm}^3$ of A, then drop P one at a time until the pink colour persists for at least 5 seconds.
3. The number of drops is recorded as a whole number. In a representative run, this might be 16 drops (which would correspond to a CuSO$_4$ concentration between 0.25% and 0.5%, used in (a)(vi)).

Key Takeaways

• The procedure is the same as for the standard concentrations; only the source of CuSO$_4$ changes.
• The recorded value is a single whole number, used as the calibration reading in (a)(vi).

Common Mistakes

• Recording a non-integer (e.g. 16.5 drops) when the procedure is drop-by-drop.
• Forgetting to record the value at all.
• Mixing up the test-tubes and recording the value for the wrong concentration.

Things to Be Careful About

• The value must be a whole number (or 'more than 30') to be credited.
• The actual number depends on the concentration of CuSO$_4$ in R and the candidate's experimental execution; the mark scheme accepts any whole number.

Background Concept

The procedure in (a)(ii) effectively produces a calibration set: each known CuSO$_4$ concentration gives a corresponding number of drops of P. To estimate the concentration of CuSO$_4$ in an unknown sample (R), the candidate's drop count for R is compared with the calibration set. If the count lies between two known values, the concentration is estimated by linear interpolation.

Understanding the Question

The candidate has recorded a number of drops for R in (a)(v) and now needs to use the results in (a)(ii) to estimate the concentration of CuSO$_4$ in R. The mark scheme requires that the estimate be based on the candidate's own results (no fixed answer is given).

Approach

Find the row in (a)(ii) whose drop count is closest to the candidate's value for R. If the value falls between two rows, interpolate linearly between the two nearest concentrations.

Step-by-Step Reasoning

1. Take the recorded number of drops for R from (a)(v). In the representative example, this is 16 drops.
2. Look up 16 in the table from (a)(ii). It lies between 14 drops (at 0.25% CuSO$_4$) and 19 drops (at 0.5% CuSO$_4$).
3. Interpolate: the fraction of the way from 0.25% to 0.5% is $\frac{16 - 14}{19 - 14} = \frac{2}{5} = 0.4$. So the estimated concentration is $0.25 + 0.4 \times (0.5 - 0.25) = 0.25 + 0.10 = 0.35\%$.
4. If the recorded value for R matches a row in the table exactly, the corresponding concentration is the answer (e.g. 14 drops → 0.25%; 19 drops → 0.5%).

Key Takeaways

• A calibration set allows the estimation of an unknown by direct comparison or interpolation.
• The estimate is only as accurate as the underlying data; large uncertainties in the drop counts translate into large uncertainties in the estimated concentration.

Common Mistakes

• Not using the candidate's own results from (a)(ii) (e.g. using a textbook value instead).
• Estimating without interpolation when the value falls between two known concentrations.
• Stating a concentration with too many significant figures (the drop counts are whole numbers, so the estimate is at best to one or two significant figures).

Things to Be Careful About

• The estimate depends on the candidate's own recorded value in (a)(v); any reasonable concentration that corresponds to that value is accepted.
• The estimate should be expressed as a percentage (e.g. 0.35%) to be consistent with the calibration data.

Background Concept

A line graph is used to display how a dependent variable (here, percentage inhibition of sucrase) changes with an independent variable (concentration of inhibitor). The independent variable is conventionally placed on the x-axis and the dependent on the y-axis. CIE conventions require:

• A scale that uses at least half the grid in both directions.
• Axis labels that include both the quantity and the unit.
• Points plotted as small dots in circles or crosses (not large blobs) so that the actual point is unambiguous.
• Points joined by a thin line (ruled straight or smooth curve), not by a hand-drawn jagged line.

Understanding the Question

The candidate is asked to plot the data in Table 1.3 on the grid in Fig. 1.3. The data are five (concentration, percentage inhibition) pairs. The expected shape of the curve is a steep rise at low concentrations followed by a plateau as the inhibitor saturates its effect (a typical dose–response curve).

Approach

Choose the x-axis scale to fit the data (0.5 to 2.5 $\text{mg cm}^{-3}$, marked every 0.5) and the y-axis scale to fit the percentage inhibition values (0 to 60%, marked every 10). Plot each of the five points carefully, then join them with a smooth curve that reflects the trend (steep rise, then plateau).

Step-by-Step Reasoning

1. The x-axis range is 0.5 to 2.5 $\text{mg cm}^{-3}$. The grid is large enough to accommodate this scale.
2. The y-axis range is 0 to 60%. The maximum data value is 52.5%, so 60% gives a small bit of headroom.
3. Mark every 0.5 on the x-axis and every 10 on the y-axis. This is the minimum density required by the mark scheme ('labelled at least every 2 cm' of grid).
4. Plot each of the five points as a small dot in a circle or a small cross:
   • (0.50, 8.0)
   • (1.00, 27.5)
   • (1.50, 44.5)
   • (2.00, 51.0)
   • (2.50, 52.5)
5. Join the points with a thin, smooth curve. The curve should rise steeply at first (between 0.5 and 1.5 $\text{mg cm}^{-3}$) and then flatten as it approaches ~53% (between 1.5 and 2.5 $\text{mg cm}^{-3}$). The point at (2.50, 52.5) is near the top of the plateau.
6. Use a sharp pencil so that points and lines are precise and easy to read.

Key Takeaways

• A dose–response curve typically shows a steep rise followed by a plateau as the system approaches saturation.
• Graph conventions: small dots/crosses, thin lines, axes labelled with quantity and unit, scales that use at least half the grid.
• The independent variable (concentration) is on the x-axis; the dependent variable (% inhibition) is on the y-axis.

Common Mistakes

• Labelling the y-axis 'percentage inhibition' without the % sign in the unit.
• Using an awkward scale (e.g. 0.3, 0.6, 0.9, 1.2...) that does not start at 0.5 or does not use half the grid.
• Plotting points as large dots or filled circles, which obscure the exact location.
• Joining the points with straight ruled lines or a hand-drawn jagged line instead of a smooth curve.
• Plotting % inhibition on the x-axis by mistake (the dependent variable goes on the y-axis).

Things to Be Careful About

• The data show a curve, not a straight line: the points must be joined with a smooth curve, not ruled straight lines.
• The y-axis scale should be chosen so the highest data point (52.5%) is well within the grid; 60% is appropriate.
• All five points must be plotted; missing a point loses a mark.

Background Concept

To find an x-value on a graph that corresponds to a given y-value (or vice versa), the standard technique is to draw construction lines. A horizontal line from the y-axis to the curve, followed by a vertical line from the curve to the x-axis, identifies the required (x, y) pair. This works for any smooth curve and is the standard way to interpolate between data points.

Understanding the Question

The candidate has plotted a smooth curve in (b)(i) and now needs to find the concentration of inhibitor that gives exactly 24% inhibition of sucrase. The mark scheme requires that two lines be drawn on the graph: one across from the y-axis, and one down to the x-axis. Both intercepts must be shown.

Approach

Draw a horizontal line from 24% on the y-axis to the curve, then a vertical line from the intersection on the curve down to the x-axis. Read off the x-value.

Step-by-Step Reasoning

1. Locate 24% on the y-axis (between the 20 and 30 gridlines, just below halfway).
2. Using a ruler, draw a horizontal dashed line from this point across the graph to the right, until it meets the curve.
3. From the point of intersection on the curve, draw a vertical dashed line down to the x-axis.
4. Read off the x-value where this vertical line meets the x-axis. Using the plotted data, this value lies between 0.5 (where inhibition is 8.0%) and 1.0 (where inhibition is 27.5%). Linear interpolation gives approximately $\frac{24 - 8.0}{27.5 - 8.0} = \frac{16}{19.5} \approx 0.82$ of the way from 0.5 to 1.0, i.e. about $0.5 + 0.82 \times 0.5 = 0.91\ \text{mg cm}^{-3}$.
5. The exact value depends on the candidate's plotted curve; the mark scheme only requires that the two construction lines be drawn clearly.

Key Takeaways

• Construction lines (horizontal then vertical) are the standard way to read an x-value from a y-value on a graph.
• The lines should be drawn with a ruler and be clearly visible (dashed lines are conventional).
• The mark scheme credits the lines, not the specific numerical value (which depends on the candidate's plotting).

Common Mistakes

• Drawing the horizontal line in the wrong direction (e.g. starting from the x-axis instead of the y-axis).
• Drawing only one of the two lines (e.g. only the horizontal, leaving the x-value unmarked).
• Drawing the vertical line down to the x-axis but failing to read off the value clearly.
• Drawing the lines freehand (without a ruler) so they are wobbly and the reading is ambiguous.

Things to Be Careful About

• The horizontal line must reach the curve, not stop at the nearest data point (24% does not coincide with any data point, so the line must intersect the curve between two points).
• The vertical line must drop from the point on the curve, not from the nearest data point.
• The lines should be drawn lightly (e.g. dashed) so they do not obscure the plotted curve.

Background Concept

Enzymes are biological catalysts with a specific 3D shape, including an active site where the substrate binds. The induced-fit model states that the active site changes shape slightly when the substrate binds, but the resting shape is determined by the enzyme's tertiary structure.

Inhibitors reduce enzyme activity by interfering with this process:

• Competitive inhibitors bind to the active site, competing with the substrate. They directly block the substrate from binding.
• Non-competitive (allosteric) inhibitors bind to a site elsewhere on the enzyme (an allosteric site), changing the shape of the active site so that the substrate can no longer bind effectively.

In both cases, the number of enzyme–substrate complexes that can form per unit time is reduced, and so the rate of reaction falls.

Understanding the Question

The candidate is asked to suggest how the inhibitor from green tea reduces the activity of sucrase. The mark scheme awards up to 2 marks for combining one point from each of two categories:

• Where/how the inhibitor binds (active site or allosteric site)
• The effect on the active site shape and substrate binding

The expected answer combines a binding statement with a consequence statement.

Approach

State that the inhibitor binds (at the active site or at an allosteric site) and that this changes the active site shape, preventing substrate binding and reducing the number of enzyme–substrate complexes.

Step-by-Step Reasoning

1. The inhibitor molecule has a shape that is complementary to a region of the sucrase enzyme.
2. The inhibitor binds either at the active site (competitive) or at a different (allosteric) site on the enzyme.
3. Binding of the inhibitor changes the 3D shape of the active site (in the case of allosteric binding) or directly occupies it (in the case of active-site binding).
4. Because the active site no longer fits the substrate (sucrose), fewer enzyme–substrate complexes can form.
5. With fewer enzyme–substrate complexes, the rate of hydrolysis of sucrose is reduced, so the activity of sucrase appears lower.

Key Takeaways

• Enzyme inhibitors work by binding to the enzyme (at the active site or an allosteric site) and either blocking the substrate or changing the active site shape.
• The result is a reduction in the number of enzyme–substrate complexes, and therefore a reduced rate of reaction.
• The mark scheme requires both a binding statement AND a consequence statement for full marks.

Common Mistakes

• Vague statements: 'the inhibitor stops the enzyme from working' — does not specify where it binds or how it changes the enzyme.
• Missing the consequence: only stating where the inhibitor binds without saying what happens to substrate binding.
• Confusing the inhibitor with the substrate (e.g. saying the inhibitor is broken down by the enzyme).
• Forgetting to mention that the number of enzyme–substrate complexes is reduced.

Things to Be Careful About

• The question asks for a 'suggestion', so the candidate is not required to commit to whether the inhibition is competitive or non-competitive. Either is acceptable as long as the binding and consequence are stated.
• The key biological terms — 'active site', 'allosteric site', 'enzyme–substrate complex' — should be used precisely.
• The mark scheme awards marks for combining one point from each category; a single point from only one category scores only 1 mark.

Background Concept

Benedict's reagent (a blue solution containing copper(II) sulfate in an alkaline citrate buffer) reacts with reducing sugars on heating. Cu²⁺ ions are reduced to Cu⁺, which precipitate as red copper(I) oxide. The colour changes from blue to green (low concentration of reducing sugar) to yellow to orange to brick red (high concentration).

To convert this qualitative colour change into a QUANTITATIVE measurement of concentration, a calibration series is needed: a set of standard solutions of known concentration is tested in exactly the same way, and the colour (or the time taken to reach a particular colour) of the unknown sample is compared with the standards.

Understanding the Question

The scientists want to know the concentration of reducing sugars in a solution after 5 minutes of sucrase activity. The candidate is asked to describe how the concentration of reducing sugars could be determined. The mark scheme requires three points:

1. A calibration series of known concentrations must be prepared.
2. Benedict's solution must be added and the mixture heated to at least 80°C (the temperature at which the reaction proceeds at a useful rate).
3. The sample must be compared with the standards using either the time taken to reach a particular colour or the final colour.

Approach

Describe a calibration-based colorimetric procedure: prepare a series of standards, treat each standard identically to the sample (Benedict's + heat), then match the sample's behaviour to the closest standard.

Step-by-Step Reasoning

1. Prepare standards: make up a series of solutions of known reducing-sugar concentration (e.g. glucose at 0, 0.5, 1.0, 1.5, 2.0, 2.5 mg cm⁻³). These provide the calibration.
2. Treat standards with Benedict's: add a fixed volume of Benedict's reagent to each standard and heat in a water bath at ≥80°C for a fixed time. Observe the colour change (and/or the time taken to reach a particular colour).
3. Treat the sample identically: add the same volume of Benedict's reagent to the sample and heat in the same way. Record the final colour (and/or the time taken to reach the same reference colour).
4. Compare with the calibration: match the sample's colour (or time) to the standard that gives the same result. The concentration of that standard is the estimated concentration in the sample.

Either the final colour or the time taken to reach a particular colour can be used as the comparison; time is often more reproducible because it is a continuous variable.

Key Takeaways

• Benedict's test on its own is qualitative (it tells you whether a reducing sugar is present, but not how much).
• To make it quantitative, a calibration series of known concentrations is essential.
• Either final colour (a discrete comparison) or time to reach a given colour (a continuous comparison) can be used as the comparison variable.
• Heating to at least 80°C is required for the Benedict's reaction to proceed at a useful rate; below this, the colour change is very slow.

Common Mistakes

• Describing only the qualitative Benedict's test ('add Benedict's, heat, observe the colour change') without mentioning a calibration series.
• Omitting the heating step or specifying an incorrect temperature (e.g. 37°C, which is body temperature and too low).
• Saying 'compare the colour' without specifying what the colour is being compared to (i.e. without mentioning the standards of known concentration).
• Forgetting to use the same volume of Benedict's and the same heating time for the standards and the sample.

Things to Be Careful About

• The comparison must be to STANDARDS of KNOWN concentration — a bare 'compare the colour' is not enough.
• The temperature threshold (≥80°C) is part of the mark scheme; omitting it loses a mark.
• The Benedict's test is for REDUCING sugars, so the procedure is specific to reducing sugars (not all sugars are reducing — sucrose is not, but its hydrolysis products glucose and fructose are).

Background Concept

A plan diagram is a low-magnification outline of a specimen that records the shapes, sizes and positions of the different tissues, without showing any individual cells. It is drawn from observation through a microscope and is used to record tissue organisation. Conventions are strict because the diagram is a scientific record: lines must be sharp and continuous, no shading is allowed, and no individual cells may be drawn inside the tissue regions. The number of lines used to represent a single-layered tissue such as an epidermis depends on the orientation of the cut: where the epidermis is cut transversely, two adjacent wall layers are seen as two lines; where a thicker cut is visible, three lines may represent the row of cells plus their bounding walls.

Understanding the Question

You are looking down the microscope at slide M1, a stained transverse section through a leaf. The shaded area in Fig. 2.1 shows the central midrib region that you must draw. You must use a sharp pencil, fill the available space, follow plan-diagram conventions, and add ONE ruled label line to the lower epidermis.

The mark scheme rewards five specific features: using most of the space and drawing the upper epidermis as three lines and the lower epidermis as two; drawing the correct region with no individual cells; the correct proportions of the vascular bundle; subdividing the vascular bundle (to show xylem and phloem); and one correctly placed label on the lower epidermis.

Approach

First, identify the upper epidermis, the lower epidermis, the mesophyll and the vascular bundle. Then plan the size of your drawing so that it fills most of the space (and is large enough to show the bundle subdivisions clearly). Draw the outlines, then check proportions against the slide, and finally add one ruled label.

Step-by-Step Reasoning

1. Look at the slide under low power and locate the shaded region from Fig. 2.1. This is the central midrib, with a downward protrusion of the lower epidermis and a flat upper surface.
2. Outline the upper epidermis as three lines. The third line represents the cell-wall boundary on the inner side of the cell row; the central line is the shared wall between adjacent epidermal cells.
3. Outline the lower epidermis as two lines where the cut passes through the protruding midrib, because only the upper and lower walls of the cells are seen in this region.
4. Leave the mesophyll blank — no palisade or spongy mesophyll cells. This is a plan, not a high-power drawing.
5. Within the midrib, draw the vascular bundle at the correct size relative to the surrounding tissues. The bundle should be large because it is the main vascular supply of the midrib.
6. Subdivide the bundle: xylem towards the upper side, phloem towards the lower side, separated by a thin line representing the cambium/bundle boundary. This earns the "subdivisions" mark.
7. Use a sharp pencil; keep all lines thin, continuous and free of shading.
8. Add ONE ruled label line that ends exactly on the lower epidermis. Write the label "lower epidermis".

Key Takeaways

• A plan diagram records tissues, not cells: no shading, no internal cell detail.
• The number of lines used to depict a single-cell layer reflects how the section is cut (two lines through the cell, three lines across a row).
• The vascular bundle must be drawn at the correct proportion and must be subdivided.
• Exactly one ruled label line is required; it must touch the structure being labelled.

Common Mistakes

• Drawing individual palisade or spongy mesophyll cells inside the plan (a high-power drawing error, not a plan error).
• Shading the vascular tissue to make it look "solid".
• Drawing the upper and lower epidermis as the same number of lines without checking the section.
• Forgetting to subdivide the bundle, so it appears as one undifferentiated mass.
• Adding more than one label, or using a label that does not end on the structure.
• Using a thick or broken line for any part of the outline.

Things to Be Careful About

The shaded region in Fig. 2.1 is the central midrib, NOT the whole leaf. Only the region indicated should be drawn. Use the available space — a small diagram wastes easy marks. The vascular bundle should look proportionate to the midrib width; an undersized bundle suggests the wrong specimen was examined.

Background Concept

A high-power (cellular) drawing shows individual cells with their walls and any visible contents. Unlike a plan diagram, individual cells ARE drawn. For a plant cell, the cell wall is shown as TWO close parallel lines (one for each side of the shared wall), and the lines must be continued all the way around the cell. Xylem vessel elements are dead, hollow cells; in transverse section they appear as roughly circular or polygonal outlines with thickened walls and an empty (clear) lumen. They are joined end-to-end, and their end walls are perforated or absent to form continuous vessels.

Understanding the Question

You are looking at slide M1 through the microscope at high power. You must select a line of FOUR adjacent xylem vessel elements (each one must touch at least one other in the line) and draw them large and accurately. You must add ONE ruled label line that ends on the wall of ONE xylem vessel element.

The mark scheme rewards: clean sharp lines with no shading; four correctly selected cells each touching at least one neighbour; a double-line wall drawn around every cell; the correct shape of xylem vessel elements; and one correct label on a cell wall.

Approach

First locate the xylem in the vascular bundle on M1. Choose a row of four elements where each one clearly touches the next. Plan the size of the drawing so the four cells fill most of the available space. Draw each cell with a double-line wall, leave the lumen empty (no contents drawn), and add one label.

Step-by-Step Reasoning

1. Centre the xylem in the field of view and choose a clean line of four xylem vessel elements. They should form a single row, each touching the next along a shared wall.
2. Plan the size of the drawing so the four cells together fill most of the available space, but cells should not overlap or run off the page.
3. Draw the outline of the first cell as two close parallel lines, then continue around the whole cell so the wall is a complete double line.
4. Where the second cell touches the first, the double-line wall of the first cell is also the double-line wall of the second; do not draw extra lines for the shared wall.
5. Repeat for the third and fourth cells, so all four cells have a complete double-line wall and they all touch.
6. Xylem vessel elements have a roughly polygonal/rounded outline with thickened walls and an empty lumen — do not shade the wall or fill the lumen.
7. Add ONE ruled label line that ends exactly on the wall of one xylem vessel element. Write the label "cell wall".
8. Check: lines continuous and thin, no shading, four cells each touching a neighbour, double-line wall around each, correct shape, one correct label.

Key Takeaways

• A cell drawing is fundamentally different from a plan diagram: cells ARE drawn, but the contents (here, lumen) are not shaded.
• Every cell wall in a cell drawing is shown as TWO close parallel lines, all the way around the cell.
• The chosen cells must be a connected line of four, each touching at least one neighbour — isolated cells do not earn the mark.
• Only one label is required and it must end on the structure named.

Common Mistakes

• Drawing single-line walls (this is a plan-diagram error carried over).
• Shading the wall or filling the lumen (xylem vessel elements are empty when mature).
• Drawing the four cells but not in a single line, or with one cell not touching a neighbour.
• Adding multiple labels, or writing a label that does not end on a wall.
• Drawing cells that are too small, or letting them overlap or run off the page.

Things to Be Careful About

The label line must be a straight ruled line that ends ON the wall, not floating in the lumen or pointing into the empty space. The drawing must be done at high power so the cells are large enough to show the wall as two lines clearly. Do not include companion cells, parenchyma or any other cell type in the line; only xylem vessel elements count for this mark.