Respirometers and Practical Investigations

This lesson is mapped to AQA 7402 Section 3.5.1 — Respiration: practical investigations and is the anchor lesson for both Required Practical 7 (investigating dehydrogenase activity in chloroplasts — the Hill reaction) and Required Practical 8 (investigating the effect of environmental variables on the rate of respiration, frequently using respirometers) (refer to the official AQA specification document for exact wording). The two practicals form a complementary pair: RP7 dissects photosynthesis by isolating the light-dependent electron-transfer step using an artificial acceptor (DCPIP); RP8 dissects respiration by measuring whole-organism gas exchange. Both train the same core analytical skills — controlled variables, replication, calibration, statistical testing, and evaluation.

Respirometry is the quantitative measurement of an organism's rate of respiration. Since aerobic respiration consumes O₂ and produces CO₂, the rate can be estimated by measuring either gas exchange. A respirometer is the apparatus used. Understanding how to set one up, run it correctly, analyse the data, and evaluate sources of error is a high-frequency exam topic — frequently worth 6–9 marks on AQA Paper 2 and 3.

Key Definition: A respirometer is a device used to measure the rate of respiration of an organism by detecting changes in gas volume — usually O₂ consumption when a CO₂ absorbent is present, or the difference between O₂ uptake and CO₂ production when no absorbent is present.

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Principles of Respirometry

Why Measure O₂ Uptake?

• In aerobic respiration, oxygen is the terminal electron acceptor in the inner-membrane ETC (lesson 3). One molecule of O₂ is consumed for every two NADH (or one NADH + one FADH₂) re-oxidised — so O₂ consumption is a direct proxy for ETC flux.
• Measuring O₂ consumption provides a quantitative, sensitive, and reliable estimate of the rate of aerobic respiration.

The Problem of CO₂ Production

• Respiration also produces CO₂, which would increase the gas volume inside a sealed chamber. This rise in CO₂ would mask the fall in volume caused by O₂ consumption — confounding the measurement.
• The solution is to place a CO₂ absorbent inside the chamber:
  • Soda lime — a granular mixture of NaOH and Ca(OH)₂ that absorbs CO₂ by forming sodium and calcium carbonates.
  • Potassium hydroxide (KOH) solution — a concentrated alkali that absorbs CO₂ by forming K₂CO₃ in solution.
• The absorbent removes CO₂ as fast as it is produced, so any change in gas volume reflects only O₂ consumption.

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Simple Respirometer Design

A typical simple respirometer consists of five components:

1. A sealed glass tube or flask containing the living organisms (e.g. germinating seeds, woodlice, maggots, or a small invertebrate). The tube is fitted with a bung carrying the manometer tube and (where appropriate) a syringe.
2. A layer of CO₂ absorbent (soda lime granules or KOH solution on cotton wool) placed near the organisms but not in direct contact (KOH is caustic and would harm living tissue).
3. A capillary manometer tube connected to the sealed chamber, containing a coloured fluid (oil or water with dye) that forms a meniscus or a discrete bubble. As O₂ is consumed and CO₂ is absorbed, the total gas volume falls, and the fluid moves along the tube towards the organism chamber.
4. A graduated scale alongside the capillary tube to measure the distance moved by the fluid over a defined time interval.
5. A reset syringe allowing the fluid to be reset to its starting position between readings without dismantling the apparatus.

How It Works

1. Organisms respire aerobically, consuming O₂ and releasing CO₂.
2. CO₂ is absorbed by the soda lime (or KOH).
3. With CO₂ removed and O₂ consumed, total gas volume in the sealed chamber decreases.
4. The decrease in volume reduces internal pressure relative to atmospheric.
5. The coloured fluid in the manometer moves towards the organism chamber until pressure equalises.
6. Rate of fluid movement (distance/time) is proportional to rate of O₂ consumption — and hence rate of respiration.

graph LR
    A["Organisms<br/>(germinating seeds<br/>or invertebrates)"] -->|"consume O₂<br/>release CO₂"| B["Chamber gas"]
    C["Soda lime / KOH"] -.->|"absorbs CO₂"| B
    B -->|"net volume falls"| D["Internal pressure drops"]
    D -->|"coloured fluid moves<br/>towards organisms"| E["Manometer reading"]
    E -->|"distance / time × π r²"| F["Rate of O₂ consumption<br/>(mm³ min⁻¹)"]
    F -->|"÷ mass"| G["Rate per unit mass<br/>(mm³ min⁻¹ g⁻¹)"]

    style A fill:#3498db,color:#fff
    style F fill:#27ae60,color:#fff
    style G fill:#e67e22,color:#fff

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Control Experiment

A control tube is set up identically to the experimental tube but with dead organisms (e.g. boiled seeds) or glass beads of equivalent volume instead of living organisms. The control:

• Accounts for any changes in gas volume caused by temperature or pressure fluctuations in the laboratory (not caused by respiration).
• Detects leaks in the apparatus.
• Ensures that any movement of fluid in the experimental tube is due solely to respiration of the living organisms.
• Provides a baseline: subtract the control reading from the experimental reading to give the true rate of O₂ consumption.

Exam Tip: Always describe the control in respirometer experiments. Without it, you cannot be confident that changes in gas volume are due to respiration rather than environmental factors. This is the single most common control-related mark loss on respirometry questions.

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Calculating Rate of Respiration

From Fluid Movement

• Measure the distance moved by the fluid in the capillary tube over a known time interval.

• If the internal radius (and hence cross-sectional area) of the capillary tube is known, the volume of O₂ consumed can be calculated:

  Volume = π × r² × distance moved

  where r = radius of the capillary tube.

• Rate of O₂ consumption = volume of O₂ consumed ÷ time.

Units

• Rate may be expressed as:
  • mm³ min⁻¹ (volume per unit time).
  • mm³ min⁻¹ g⁻¹ (volume per unit time per unit mass — to allow fair comparison between organisms of different sizes or between batches of differing mass).
  • cm³ h⁻¹ kg⁻¹ for larger animals.

Exam Tip: When comparing respiration rates between different organisms or different conditions, always express the rate per unit mass. Otherwise a large organism trivially appears to "respire faster" than a small one, when in fact what differs is mass, not mass-specific metabolic rate.

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Calculating the Respiratory Quotient (RQ)

As covered in lesson 4, the RQ is:

RQ = CO₂ produced ÷ O₂ consumed

Using a Respirometer to Measure RQ

To determine RQ, take two paired measurements:

1. With soda lime (CO₂ absorbed): The change in gas volume equals the volume of O₂ consumed (because the CO₂ produced is removed instantly).
2. Without soda lime (CO₂ NOT absorbed): The change in gas volume equals (O₂ consumed) − (CO₂ produced), because the released CO₂ partially replaces the volume lost to O₂ consumption.

• O₂ consumed = volume change with soda lime.
• CO₂ produced = O₂ consumed − volume change without soda lime.
• RQ = CO₂ produced ÷ O₂ consumed.

Worked Example

Condition	Volume change (mm³ per minute)
With soda lime	0.8 mm³ min⁻¹ decrease
Without soda lime	0.2 mm³ min⁻¹ decrease

• O₂ consumed = 0.8 mm³ min⁻¹.
• Net volume change without soda lime = 0.2 mm³ min⁻¹ = (O₂ consumed) − (CO₂ produced).
• CO₂ produced = 0.8 − 0.2 = 0.6 mm³ min⁻¹.
• RQ = 0.6 ÷ 0.8 = 0.75.

This RQ of 0.75 suggests the organism is respiring a mixture of substrates, or primarily lipid (RQ ≈ 0.7 for pure lipid).

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Investigating the Effect of Temperature on Respiration Rate (RP8)

Method

1. Set up respirometers at a range of temperatures using thermostatically controlled water baths (e.g. 10 °C, 20 °C, 30 °C, 40 °C, 50 °C).
2. Allow the organisms to acclimatise at each temperature for a set time (5–10 min) before taking measurements.
3. Measure the rate of O₂ consumption at each temperature (record distance moved by fluid per minute over a 5-minute window).
4. Run a control at each temperature.
5. Repeat measurements at each temperature (≥3 replicates) and calculate a mean and standard error.

Expected Results

• Rate increases with temperature up to an optimum (~35–40 °C for many small invertebrates and many germinating-seed metabolic enzymes).
• Rate decreases sharply above the optimum (enzyme denaturation).
• Q₁₀ can be calculated as (rate at T+10) ÷ (rate at T); typical values 1.8–2.5 for respiration.

Controlled Variables

• Mass / number of organisms.
• Volume of soda lime.
• Duration of measurement.
• Type and physiological state of organisms (use the same batch).
• Equilibration time.
• Atmospheric pressure (note any large changes; ideally repeat the experiment on a stable day).

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Investigating the Effect of Substrate on Respiration Rate

Method

1. Germinate seeds with different storage substrates: carbohydrate-rich (wheat, barley, rice) vs lipid-rich (sunflower, peanut, sesame) vs protein-rich (legumes).
2. Set up respirometers with equal masses of each seed type.
3. Measure O₂ consumption rate and CO₂ production (the paired RQ method) and calculate RQ.

Expected Results

• Carbohydrate-rich seeds: RQ ≈ 1.0.
• Lipid-rich seeds: RQ ≈ 0.7.
• Protein-rich seeds: RQ ≈ 0.8.

This is a textbook example of how the same experimental apparatus can reveal different aspects of metabolism depending on what is varied.

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Required Practical 7: Investigating Dehydrogenase Activity in Chloroplasts (The Hill Reaction)

RP7 sits within this respirometer-and-practical-skills lesson because, although it concerns photosynthesis specifically, it uses an analogous experimental logic to a respirometer — measuring an electron-transfer process by tracking a chemical change. The Hill reaction was demonstrated in 1937 by Robert Hill (paraphrased): isolated chloroplasts, when illuminated in the presence of an artificial electron acceptor (such as ferricyanide or, in modern A-Level practice, DCPIP), reduce the acceptor and release oxygen. This proved that water-splitting and electron transfer can occur in the absence of CO₂ fixation — confirming that photosynthesis is mechanistically two separable stages.

DCPIP as an Artificial Electron Acceptor

DCPIP (2,6-dichlorophenolindophenol) is a redox dye:

• Oxidised form: blue.
• Reduced form: colourless.

When isolated chloroplasts are illuminated with DCPIP, DCPIP intercepts electrons from PSI (in place of NADP⁺), is reduced, and turns colourless. The rate at which DCPIP loses colour — measured with a colorimeter — is proportional to the rate of dehydrogenase activity (i.e. the rate of the light reactions).

Method

1. Grind fresh leaf tissue (typically spinach or lettuce) with ice-cold isolation buffer (containing sucrose to maintain osmotic balance and phosphate buffer to maintain pH).
2. Filter through muslin to remove debris.
3. Centrifuge briefly to pellet chloroplasts.
4. Resuspend in fresh buffer to give a uniform chloroplast suspension.
5. In a test tube, mix the chloroplast suspension with DCPIP solution.
6. Place at a measured distance from a light source.
7. At fixed time intervals (e.g. every 30 s for 5 min), measure absorbance at 605 nm using a colorimeter.
8. Run a control in the dark (no light → no electron transfer → no DCPIP reduction).
9. Run a boiled-chloroplast control (heat-denatured enzymes → no dehydrogenase activity).

Variables

• Independent variable: e.g. light intensity, temperature, or chloroplast concentration.
• Dependent variable: rate of DCPIP decolourisation (change in absorbance per unit time).
• Controlled variables: chloroplast concentration, DCPIP concentration, buffer pH and ionic strength, temperature (water bath), wavelength of light.

Expected Results

• In illuminated tubes with active chloroplasts, absorbance falls progressively as DCPIP is reduced.
• In the dark control, absorbance remains constant (no dehydrogenase activity without light to drive electron transport).
• In the boiled-chloroplast control, absorbance remains constant (denatured enzymes cannot drive electron transport).
• Rate of decolourisation increases with light intensity, up to a saturation point (consistent with the limiting-factor model in lesson 7).

Sources of Error and Improvements