High School Reading: Grades 11-12 Quiz: Follow Complex Multistep Procedures In Technical Texts

Protocol: Isolation of Genomic DNA from Plant Leaf Tissue (CTAB Method)

1. Grind 100 mg of fresh leaf tissue to a fine powder under liquid nitrogen using a chilled mortar and pestle. Do not allow the tissue to thaw at any point during grinding.

2. Transfer the frozen powder to a 2 mL microcentrifuge tube containing 700 μL of preheated (65 °C) CTAB extraction buffer.

3. Invert the tube several times, then incubate in a water bath at 65 °C for 30 minutes, inverting every 10 minutes.

4. Allow the tube to cool to room temperature for 5 minutes. Then add 700 μL of chloroform:isoamyl alcohol (24:1) and mix by gentle inversion for 5 minutes. Vigorous shaking will shear the DNA.

5. Centrifuge at 12,000 × g for 10 minutes at 4 °C. Three layers will form.

6. Transfer only the upper aqueous phase to a fresh tube, avoiding the interface. Repeat steps 4–6 if the interface is heavily pigmented.

7. Precipitate DNA by adding 0.7 volumes of ice-cold isopropanol; invert gently until strands become visible.

8. Pellet the DNA at 12,000 × g for 5 minutes, wash the pellet twice with 70% ethanol, air-dry, and resuspend in 50 μL TE buffer.

A researcher's final DNA yield is normal by mass, but agarose gel electrophoresis shows a bright smear of low-molecular-weight fragments rather than a tight high-molecular-weight band. Which departure from the protocol best accounts for this specific pattern of results?

Procedure: Preparing a Standard Curve for Protein Quantification (Bradford Assay)

1. Prepare a 1.00 mg/mL bovine serum albumin (BSA) stock solution in the same buffer used for the unknown samples.

2. In a rack of labeled tubes, prepare 500 μL standards spanning 0, 2, 4, 8, 12, 16, and 20 μg BSA by diluting the stock with buffer. Prepare in duplicate.

3. To each tube, add 500 μL of Bradford reagent. Vortex briefly and incubate at room temperature for exactly 5 minutes.

4. Between 5 and 20 minutes after adding reagent, transfer each solution to a disposable cuvette and read absorbance at 595 nm, blanking the spectrophotometer against the 0 μg standard.

5. Plot mean absorbance versus micrograms of BSA. Fit a linear regression only to the linear portion of the curve; the assay typically loses linearity above ~15 μg protein.

6. For each unknown, dilute if necessary so that the reading falls within the linear range, then interpolate protein content from the fitted equation. Multiply by the dilution factor to obtain protein in the original sample.

A student's unknown sample gives an absorbance of 0.612. Her linear regression, fit through all seven standards, yields $A = 0.030x + 0.010$, and she reports 20.1 μg protein. A supervisor insists the reported value is systematically wrong. What is the most likely source of the error?

Procedure: Determining the Equivalence Point of a Weak-Acid Titration

1. Pipette exactly 25.00 mL of the unknown weak-acid solution into a clean 250 mL Erlenmeyer flask. Add ~50 mL of distilled water and a stir bar.

2. Rinse and fill a 50 mL burette with standardized 0.1000 M NaOH. Record the initial burette reading to the nearest 0.02 mL.

3. Insert a calibrated pH probe into the flask. Begin gentle stirring, ensuring the probe does not contact the stir bar.

4. Add NaOH in 1.00 mL increments, recording pH after each addition once the reading stabilizes for ≥10 s.

5. When pH begins to change by more than 0.3 units per addition, reduce the increment to 0.20 mL. Continue until pH has passed clearly beyond 11.

6. Plot pH versus volume added. Locate the equivalence point at the midpoint of the steepest rise on the curve (the inflection point).

7. Use the volume $V_{eq}$ at the equivalence point and the titrant concentration to compute the moles of acid, and thus its original molarity.

8. To estimate $pK_a$, read the pH at the half-equivalence volume $V_{eq}/2$; this pH equals $pK_a$.

A student's titration curve shows a clear inflection at 22.40 mL, but she mistakenly identifies the equivalence point as the volume where pH first equals 7.00, at 18.00 mL. If she then reports $pK_a$ using her (incorrect) $V_{eq}$, how will her reported $pK_a$ compare with the true value?

Procedure: Gram Staining of a Bacterial Smear

1. Using an inoculating loop, transfer a small amount of bacterial culture to a drop of distilled water on a clean glass slide. Spread thinly.

2. Air-dry the smear completely, then heat-fix by passing the slide (smear-side up) through a Bunsen flame 2–3 times. Overheating will distort cell morphology.

3. Flood the smear with crystal violet for exactly 60 seconds. Rinse briefly with distilled water.

4. Flood with Gram's iodine mordant for exactly 60 seconds. Rinse briefly.

5. Decolorize with 95% ethanol drop by drop, tilting the slide, until runoff is nearly clear. This step is critical and typically requires 3–10 seconds; over-decolorization will strip stain from gram-positive cells.

6. Rinse immediately with distilled water to arrest decolorization.

7. Counterstain with safranin for 60 seconds. Rinse, blot dry, and examine under oil immersion.

8. Gram-positive cells appear purple; gram-negative cells appear pink.

A student stains a known Staphylococcus aureus culture (a gram-positive organism) but consistently observes pink cocci under the microscope. Which single deviation from the procedure most specifically produces this result while leaving cell morphology intact?

Field Procedure: Estimating Stream Discharge by the Float Method

1. Select a straight reach of stream at least 10 m long with uniform width and minimal turbulence. Mark an upstream point A and a downstream point B exactly 10.0 m apart.

2. At three evenly spaced cross-sections between A and B, measure stream width and record depth at intervals of one-fifth the width. Average all depth readings and multiply by the average width to obtain the mean cross-sectional area, $A_c$ (in m²).

3. Release a partially submerged float (e.g., an orange) upstream of A. Using a stopwatch, record the time $t$ (in seconds) for the float to travel from A to B. Repeat five times and take the mean.

4. Compute the surface velocity: $v_s = 10.0 / t$ (m/s).

5. Because surface velocity exceeds mean velocity, multiply by a correction coefficient $k$: use $k = 0.85$ for smooth beds, $k = 0.75$ for rocky or weedy beds. This gives mean velocity $v_m = k \cdot v_s$.

6. Discharge $Q$ in m³/s is then $Q = A_c \cdot v_m$.

A student measures a rocky-bedded stream and obtains an average cross-sectional area of $1.20\ \text{m}^2$ and a mean float time of 25.0 s. What is the correctly computed discharge?

Procedure: Calibrating a Pipette by Gravimetric Analysis

1. Bring a bottle of distilled water to the calibration room at least 2 hours in advance so its temperature equilibrates with the ambient air. Record the water temperature $T$ to the nearest 0.1 °C.

2. Set the adjustable pipette to its nominal volume (e.g., 100.0 μL). Place a clean, dry weighing vessel with a tight-fitting lid on the analytical balance and tare.

3. Pre-wet the pipette tip by aspirating and dispensing distilled water three times before the first measurement.

4. Aspirate water and dispense it into the weighing vessel. Close the lid immediately to prevent evaporation. Record the mass $m_i$.

5. Repeat step 4 for a total of ten measurements, taring between dispenses.

6. Look up the Z-factor for the recorded temperature (e.g., $Z = 1.0029$ μL/mg at 22.0 °C). This factor corrects for water density and buoyancy.

7. Convert each mass to volume: $V_i = m_i \cdot Z$. Compute the mean $\bar{V}$ and the standard deviation $s$.

8. Report inaccuracy as $(\bar{V} - V_{\text{nominal}})/V_{\text{nominal}} \times 100\%$ and imprecision as $s/\bar{V} \times 100\%$.

A technician calibrating a 100 μL pipette obtains ten dispenses with a mean mass of 99.20 mg at 22.0 °C ($Z = 1.0029$ μL/mg). The manufacturer specifies inaccuracy must lie within ±0.8%. What is the correct conclusion?

Procedure: Preparing Recombinant Plasmid DNA by Restriction Digestion and Ligation

1. In two separate 20 μL reactions, digest 1 μg of vector DNA and 1 μg of insert DNA with the same restriction enzyme (e.g., EcoRI). Include 2 μL of the manufacturer's 10× buffer and incubate at 37 °C for 1 hour.

2. Heat-inactivate the enzyme at 65 °C for 20 minutes.

3. To prevent vector self-ligation, treat the digested vector only with 1 U of shrimp alkaline phosphatase for 30 minutes at 37 °C, then heat-inactivate at 65 °C for 15 minutes. Do not phosphatase-treat the insert.

4. Run 2 μL of each digest on a 1% agarose gel to verify complete digestion (single band of expected size for the vector; insert band visible).

5. Combine 50 ng vector and a 3:1 molar excess of insert in a 20 μL ligation reaction with T4 DNA ligase and 1× ligase buffer.

6. Incubate at 16 °C overnight, then heat-inactivate at 65 °C for 10 minutes.

7. Transform 2 μL of the ligation mixture into competent E. coli and plate on selective medium.

After transformation, a researcher observes hundreds of colonies on the vector-only control plate (ligation without insert), nearly as many as on the vector-plus-insert plate. Screening reveals that most colonies from both plates contain vector without insert. Which single deviation from the protocol most directly explains this pattern?

Procedure: Serial Dilution for Bacterial Enumeration

1. Label seven sterile tubes 10⁻¹ through 10⁻⁷. Aseptically dispense 9.0 mL of sterile saline into each tube.

2. Vortex the original bacterial culture. Transfer 1.0 mL into the 10⁻¹ tube and vortex thoroughly. This produces a 1:10 dilution of the original.

3. Using a fresh sterile pipette tip, transfer 1.0 mL from 10⁻¹ into 10⁻². Vortex. Continue serially through 10⁻⁷, changing tips at each step.

4. From each of the last four tubes (10⁻⁴ through 10⁻⁷), pipette 0.1 mL onto a labeled nutrient-agar plate and spread evenly.

5. Invert plates and incubate at 37 °C for 24 hours.

6. Count only plates containing between 30 and 300 colonies. Plates outside that range are statistically unreliable.

7. Compute colony-forming units per mL of original culture as: $\text{CFU/mL} = \frac{\text{colonies counted}}{\text{dilution factor} \times \text{volume plated (mL)}}.$

A microbiologist counts 240 colonies on the 10⁻⁶ plate (all other plates being outside the countable range). Applying step 7 precisely, what is the correctly computed CFU/mL in the original culture?