Life Cycle of Hookworm (3 marks)

Hookworms infecting humans are mainly Ancylostoma duodenale and Necator americanus. Their life cycle is completed in a single host (human) and passes through the following stages:

1. Egg stage (in soil): Adult worms live in the small intestine and lay eggs (about 10,000–20,000 eggs/day for A. duodenale). Eggs pass out with faeces into moist, warm soil. In soil, eggs hatch within 1–2 days into rhabditiform (first-stage) larvae.

2. Larval development in soil: The rhabditiform larva feeds on bacteria, moults twice, and becomes the filariform (third-stage) larva, which is the infective form. This takes about 5–10 days. Filariform larvae cannot feed and survive on stored food; they remain infective for several weeks in moist shaded soil.

3. Infection of human host: The filariform larva penetrates the intact skin (usually between the toes, hence "ground itch") when a person walks barefoot on contaminated soil. After skin penetration, larvae enter blood or lymph vessels.

4. Migration in the body: Larvae are carried to the heart and then to the lungs, where they break out of capillaries into the alveoli. They migrate up the respiratory tract to the pharynx, are swallowed, and reach the small intestine.

5. Adult stage: In the small intestine, larvae moult twice and mature into adult male and female worms. The adult worm attaches to the intestinal mucosa with its buccal capsule and sucks blood. The female lays eggs, which pass in faeces, completing the cycle. The whole cycle takes about 5–7 weeks.

Laboratory Diagnosis of Hookworm (2 marks)

1. Stool examination (microscopy): A stool sample is examined for characteristic hookworm eggs — oval/ellipsoidal, colourless, with a thin transparent shell and segmented (4–8 cell stage) contents. Direct saline wet mount and iodine mount are used. Concentration techniques such as formol-ether concentration or saturated salt flotation increase the yield when the egg load is low.

2. Stool culture (Harada–Mori filter paper culture): A stool sample is cultured on filter paper strips in a test tube with water for 7–10 days. Filariform larvae migrate into the water and are identified under the microscope. This helps to differentiate A. duodenale from N. americanus and to confirm the diagnosis.

3. Blood examination: Eosinophilia (increased eosinophil count) is commonly seen in hookworm infection and supports the diagnosis.

4. Occult blood test: A positive faecal occult blood test may be found due to intestinal bleeding caused by the worms.

Note: Egg counting by Kato–Katz technique is used to assess the intensity of infection (eggs per gram of faeces).

Gram's Stain — Principle

Gram's stain is a differential staining technique developed by Hans Christian Gram (1884) used to classify bacteria into two groups on the basis of the structure of their cell wall.

Principle:

1. Primary stain (Crystal violet): All bacteria take up crystal violet and become purple, because crystal violet penetrates the peptidoglycan layer of both gram +ve and gram −ve bacteria.
2. Mordant (Gram's iodine): Iodine combines with crystal violet to form a large crystal violet–iodine (CV–I) complex inside the cell. This complex is too large to escape through an intact cell wall, so it is trapped in the thick peptidoglycan of gram +ve bacteria.
3. Decolourisation (95% alcohol/acetone): This is the critical differentiating step.
   • In gram +ve bacteria, the thick peptidoglycan layer (with little lipid) is dehydrated by alcohol, its pores shrink, and the CV–I complex is retained → cells remain purple/violet.
   • In gram −ve bacteria, the thin peptidoglycan layer and the high lipid content of the outer membrane are dissolved by alcohol, so the CV–I complex leaches out → cells become colourless.
4. Counterstain (Safranin/dilute carbol fuchsin): The decolourised gram −ve bacteria take up the pink/red counterstain, while gram +ve bacteria remain purple (unaffected by the counterstain).

Result: Gram +ve = purple/violet; Gram −ve = pink/red.

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Difference in Morphology of Gram +ve and Gram −ve Bacteria

Note: The difference in morphology is due to the structure of the cell wall, not the shape of the cell — both groups may be cocci or bacilli.

Stool Collection, Processing and Microscopic Examination

1. Stool Collection (1.5 marks)

Principle: A fresh, uncontaminated stool sample is collected in a clean, wide-mouthed container for detection of ova, cysts, parasites, pus cells, RBCs, and undigested food particles.

Materials required: Clean dry wide-mouthed container with lid, wooden applicator stick or spatula, gloves, labels, clean glass slide, normal saline, iodine solution, cover slips, microscope.

Procedure:

1. Explain the procedure to the patient and provide a clean, dry, wide-mouthed container with a tight-fitting lid.
2. Instruct the patient to pass stool into a clean bedpan or directly into the container; avoid mixing with urine, water or toilet paper.
3. Collect an adequate amount — about the size of a walnut (5–10 g) for a formed stool, or 5–10 mL if liquid.
4. Collect from different parts of the stool, and especially from any mucus, blood or pus-containing portion.
5. Label the container with the patient's name, age, sex, hospital number, date and time of collection.
6. Send the sample to the laboratory immediately (within 30 minutes to 1 hour) to prevent degeneration of trophozoites and changes in pH.

Precautions:

• Do not collect stool mixed with urine, water or disinfectant.
• Avoid oily or barium-contrast contaminated samples.
• For amoebic trophozoites, the sample must be warm and examined within 30 minutes.
• If delay is unavoidable, preserve in 10% formalin or SAF (sodium acetate–acetic acid–formalin) solution.

2. Processing of Stool Sample (2 marks)

A. Macroscopic (Physical) Examination:

• Colour: Normally brown. Pale/clay — biliary obstruction; black/tarry — upper GI bleeding; red — lower GI bleeding; green — diarrhoea or spinach.
• Consistency: Formed, semi-formed, loose, watery, or hard.
• Odour: Normally feculent; offensive in amoebiasis; sour in carbohydrate indigestion.
• Mucus, blood, pus: Note presence and amount.
• Adult worms or segments: Look for Ascaris, Taenia segments, etc.

B. Chemical Examination:

• Occult blood test (benzidine or guaiac test) — for hidden bleeding.
• pH — using pH paper.
• Reducing sugar — for lactose intolerance.

C. Concentration Techniques (for ova and cysts):

• Direct saline and iodine wet mount — routine method.
• Formalin-ether concentration — most commonly used; ova and cysts concentrate at the bottom.
• Saturated salt flotation — for light ova (e.g., Enterobius, Taenia).
• Zinc sulphate flotation — for cysts.

3. Microscopic Examination (1.5 marks)

Procedure:

1. Place one drop of normal saline on the left half of a clean glass slide and one drop of Lugol's iodine on the right half.
2. With a wooden applicator, take a small portion of stool (about the size of a match head) and emulsify in saline to make a thin uniform suspension; similarly prepare the iodine mount.
3. Apply cover slips and examine first under low power (10×) to scan the entire field, then under high power (40×) for detailed morphology.

Findings to report:

Significance:

• Saline mount — shows motile trophozoites, ova and larvae.
• Iodine mount — stains glycogen and nuclei of cysts for identification.
• Pus cells and RBCs — indicate inflammation or invasive infection.
• Charcot-Leyden crystals — allergic or parasitic infestation.

Precautions during microscopy:

• Prepare thin mounts — thick mounts obscure parasites.
• Examine the entire slide systematically before reporting "no ova or cyst seen".
• Use a warm slide for trophozoites to keep them motile.

ESR (Erythrocyte Sedimentation Rate) is the rate at which red blood cells settle down in a vertical column of anticoagulated blood in one hour, expressed in mm in the first hour. It is a non-specific acute-phase indicator of inflammation.

Evaluation of ESR (2 marks)

Westergren method (reference/standard method):

1. Blood is collected in EDTA or mixed with 3.8% trisodium citrate in the ratio 4:1 (blood : citrate).
2. The blood is mixed well and drawn into a Westergren tube (300 mm long, 2.5 mm internal bore, graduated 0–200 mm) up to the 0 mark, without air bubbles.
3. The tube is set exactly vertical in the Westergren stand, at room temperature (18–25 °C), away from direct sunlight, draught and vibration.
4. The reading is taken exactly at the end of 1 hour as the height of clear plasma above the red cell column, in mm.
5. Normal values: men 0–15 mm/1st hr, women 0–20 mm/1st hr, children 0–10 mm/1st hr, newborns 0–2 mm/1st hr.

Wintrobe method: Wintrobe tube (100 mm long, 2.5 mm bore) is used with undiluted EDTA blood; less sensitive than Westergren. Automated/micro-ESR methods are also used in modern laboratories.

Factors affecting ESR (3 marks)

A. Physiological factors

• Age: raised in the elderly; very low in newborns and infants.
• Sex: higher in females than males (lower haematocrit, hormonal effect).
• Pregnancy: rises from the 3rd month onwards.
• Menstruation: slight rise.
• Obesity and race: higher in obese persons and in some ethnic groups.

B. Pathological factors

• Increased ESR: any condition raising plasma fibrinogen or globulins (acute-phase proteins) → increased rouleaux formation. Examples: acute and chronic infections, tuberculosis, rheumatic fever, rheumatoid arthritis, collagen diseases, multiple myeloma, nephrotic syndrome, malignant tumours, tissue necrosis/myocardial infarction, and anaemia.
• Decreased ESR: polycythaemia, sickle cell disease and other poikilocytoses, hereditary spherocytosis, hypofibrinogenaemia, severe leucocytosis, congestive heart failure, and after steroid or salicylate therapy.

C. Technical (laboratory) factors

• Temperature: high temperature falsely raises ESR; low temperature lowers it.
• Time: reading must be taken at exactly 1 hour; delay in setting or reading gives a false low value. Blood older than 2 hours gives falsely low ESR.
• Position of tube: the tube must be strictly vertical — tilting increases ESR.
• Tube factors: internal bore, length, and cleanliness (grease, dirt or moisture in the tube lowers ESR).
• Anticoagulant: excess EDTA or wrong blood-to-anticoagulant ratio lowers ESR; citrate dilution must be accurate.
• Dilution errors in the Westergren method.
• Vibration, direct sunlight and draughts during the test lower or alter the reading.
• Anaemia lowers blood viscosity and falsely raises ESR, so correction for packed cell volume (Wintrobe's correction) is applied.
• Drugs such as steroids, salicylates and quinine lower ESR.

Because so many factors influence it, ESR must always be interpreted along with the clinical picture and other laboratory findings.

Sources of errors in routine haematological tests (3 marks)

Errors in routine haematology (Hb, TC, DC, ESR, platelet count, PCV) arise mainly from three stages:

1. Pre-analytical (pre-examination) errors — most common

• Patient-related: improper posture during venipuncture (standing causes haemoconcentration), exercise, stress, smoking, dehydration, or drugs (e.g. adrenaline, diuretics) altering counts.
• Specimen collection: prolonged venous stasis from a tight tourniquet, wrong order of draw, blood drawn from an arm receiving IV fluids, or capillary blood from a cold, oedematous or cyanotic finger.
• Anticoagulant faults: wrong anticoagulant (heparin for cell counts), wrong concentration of EDTA (excess causes cell shrinkage and dilution; too little causes microclots), or use of powdered/expired EDTA.
• Transport and storage: delay beyond 2–4 hours, exposure to heat or sunlight, vigorous shaking causing haemolysis, or storage at wrong temperature.
• Labelling: misidentification of patient or sample, unlabelled or illegibly labelled tubes.

2. Analytical (examination) errors

• Pipetting and dilution: inaccurate pipettes, dirty or wet pipettes, faulty dilution (e.g. 1:20 for WBC), errors in Turk's or Hayem's fluid preparation.
• Counting: uneven distribution of cells in the chamber, improper filling (air bubbles, over/under-charging), wrong choice of chamber area, counting errors, and fatigue of the technician.
• Instrument-related: dirty or scratched glassware, uncalibrated instruments, blocked apertures in automated counters, incorrect settings, or faulty reagents (contaminated stains, deteriorated diluents).
• Staining: too thick or too thin blood film, poor fixation, over- or under-staining, and incorrect pH of buffer affecting dye uptake.
• Personal bias: observer variation in identifying cells (especially band forms, eosinophils, atypical lymphocytes).

3. Post-analytical errors

• Wrong transcription of results, calculation mistakes in absolute counts, reporting in wrong units, or reporting results of one patient under another's name.

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Ways to minimize these errors (2 marks)

Pre-analytical:

• Use clean, dry, siliconized or disposable syringes and properly labelled tubes; follow the correct order of draw.
• Use the correct anticoagulant in correct proportion — EDTA at 1.5–2.0 mg per mL of blood; mix gently by inverting 8–10 times.
• Avoid prolonged tourniquet application; collect blood with the patient seated and relaxed.
• Process the sample within 2–4 hours; store at 4 °C if delay is unavoidable; never shake vigorously.

Analytical:

• Calibrate pipettes, instruments and automated counters regularly; use only fresh, properly stored reagents and stains.
• Prepare blood films correctly (thin, well-spread, properly fixed) and use buffered stain at correct pH.
• Charge the counting chamber properly, allow cells to settle, and count in a systematic pattern; count a sufficient number of cells (at least 100 WBCs for DC, 400 cells for accuracy).
• Use quality control materials (normal and abnormal controls) daily and participate in external quality assessment (EQAS).
• Ensure technicians are well trained and rotate duties to reduce fatigue and observer bias.

Post-analytical:

• Verify all calculations and transcriptions; double-check patient identity before reporting; use standard report formats with reference ranges.

By controlling errors at each stage — especially the pre-analytical stage, which accounts for the majority of errors — the reliability and reproducibility of routine haematological results can be greatly improved.

Neubauer chamber is a thick glass slide used for counting cells under the microscope. It has a ruled central counting area etched on its surface. The ruled area is 3 mm × 3 mm and is divided into 9 large squares, each 1 mm × 1 mm. The four corner squares (WBC squares) are meant for counting white blood cells, and each is further divided into 16 small squares. A special thick cover slip is placed over the ruled area, creating a known depth of 0.1 mm. Thus the volume of one WBC square = 1 mm × 1 mm × 0.1 mm = 0.1 mm³ (µL).

Total WBC count by Neubauer chamber:

Principle: Blood is diluted in a fluid that lyses red cells and stains the nuclei of white cells. The diluted blood is filled into the chamber and WBCs are counted in the four corner squares. The count is then calculated per µL (or per litre) of blood.

Requirements:

• Neubauer chamber (improved) with cover slip
• WBC diluting fluid (Turk's fluid: glacial acetic acid 2 mL, gentian violet 1 mL, distilled water 100 mL)
• Blood sample (capillary or venous, mixed with EDTA)
• WBC pipette (Thoma pipette) or micropipette
• Microscope, gauze, lancet, spirit swab

Procedure:

1. Clean and dry the Neubauer chamber and place the cover slip over the ruled area.
2. Draw blood up to the 0.5 mark in the WBC pipette (if using Thoma pipette).
3. Wipe the tip of the pipette to remove excess blood.
4. Draw WBC diluting fluid up to the 11 mark, giving a dilution of 1:20.
5. Mix gently by rotating the pipette between the palms for 2–3 minutes. Discard the first 2–3 drops.
6. Place one drop at the edge of the cover slip; the fluid is drawn in by capillary action. Ensure the chamber is neither overfilled nor underfilled and no air bubbles are present.
7. Allow 2–3 minutes for the cells to settle.
8. Under low power (10×), locate the four corner squares. Then count under high power (40×).
9. Count all WBCs in the four corner squares, following the rule: count cells lying inside the square and on the top and left borders; do not count those on the bottom and right borders.

Calculation:

$\text{Total WBC count/µL} = \frac{\text{No. of cells counted} \times \text{dilution factor}}{\text{volume of the four squares}}$

$= \frac{N \times 20}{4 \times 0.1} = N \times 50$

where N = total number of WBCs counted in the four corner squares.

Normal value: 4,000–11,000 WBCs/µL (4–11 × 10⁹/L) of blood.

(a) Classification of helminthic parasites (5 marks)

Helminths are multicellular parasitic worms. They are divided into two phyla and three medically important classes.

(b) Diagnosis of helminthic infestation by stool microscopy (5 marks)

1. Specimen collection: collect a fresh, clean stool sample in a wide-mouthed container, free from urine, water and soil. Record colour, consistency, blood, mucus and any visible worms or proglottids (macroscopic examination).
2. Direct wet mount: emulsify a small amount of stool in a drop of normal saline on a slide, cover with a coverslip, and examine under the $10\times$ and $40\times$ objectives for eggs, larvae and motile forms. A second mount with Lugol's iodine helps show detail of cysts and eggs.
3. Concentration techniques (when eggs are scanty):
   • Flotation (saturated salt or zinc sulphate solution): eggs of lower specific gravity, such as hookworm, Ascaris and Trichuris, float to the surface and are lifted off with a coverslip or loop.
   • Sedimentation (formol-ether or formalin-ethyl acetate): eggs sediment after centrifugation; useful for heavier eggs such as flukes and for tapeworm eggs.
4. Identification: eggs or larvae are identified by size, shape, shell, colour and contents. For example, Ascaris has a mammillated shell, Trichuris a barrel shape with polar plugs, and hookworm a thin shell with a segmented embryo. Pinworm eggs are better found by the cellophane (Scotch) tape perianal swab than in stool.
5. Reporting: report the parasite and its stage (egg, larva or adult), and the egg count by Kato-Katz when intensity is needed.

Sterilization and Disinfection

Definition of Sterilization: Sterilization is the complete elimination or destruction of all viable forms of microbial life, including bacteria, viruses, fungi, spores, and prions, from a material or environment.

Definition of Disinfection: Disinfection is the process of eliminating or reducing the number of pathogenic microorganisms from inanimate objects or surfaces. It does not necessarily kill all microbial life, including spores, which is the key difference from sterilization.

Classification of Sterilization Methods: Sterilization methods can be broadly classified into physical methods and chemical methods.

1. Physical Methods of Sterilization:

• Moist Heat Sterilization (Autoclaving):
  • Explanation: This method uses pressurized steam (usually at $121^\circ\text{C}$ at 15 psi for 15–20 minutes). The high temperature and moisture penetrate materials effectively, denaturing microbial proteins and destroying spores. It is the most common method for heat-stable instruments.
• Dry Heat Sterilization:
  • Explanation: This method uses high temperatures in an oven (e.g., $160^\circ\text{C}$ to $180^\circ\text{C}$ for 1–2 hours). It is used for materials that are sensitive to moisture, such as oils, powders, and glassware.
• Filtration:
  • Explanation: This method is used to sterilize heat-sensitive liquids (like antibiotics or serums). The liquid is passed through a membrane filter with a pore size small enough (typically $0.22 \mu\text{m}$) to trap bacteria and microorganisms.
• Radiation Sterilization:
  • Explanation: This involves exposing materials to high levels of ionizing radiation, such as gamma rays or electron beams. This radiation damages the DNA of microorganisms, rendering them non-viable. It is commonly used for disposable medical equipment.

2. Chemical Methods of Sterilization (Chemical Disinfection/Sterilization):

• Ethylene Oxide (EtO) Gas Sterilization:
  • Explanation: EtO is a highly effective gaseous chemical sterilant used for heat-sensitive and moisture-sensitive equipment (like plastics). It works by alkylating the proteins and nucleic acids of the microbes. However, it is toxic and requires careful aeration.
• Chemical Agents (Liquid/Gas):
  • Explanation: Various chemical solutions are used. Examples include glutaraldehyde, hydrogen peroxide, and formaldehyde. These agents denature microbial proteins. They are often used for high-level disinfection or sterilization of endoscopes and other complex instruments.

Law of Colorimeter (Beer–Lambert Law)

A colorimeter works on the combined Beer–Lambert law, which states that when a monochromatic light passes through a coloured solution, the amount of light absorbed is directly proportional to the concentration of the coloured substance and the path length (thickness) of the solution.

Statement of the law:

$A = \varepsilon \, c \, l$

Where:

• A = absorbance (optical density) = log (I₀ / Iₜ)
• ε = molar absorptivity (a constant for the substance at a given wavelength)
• c = concentration of the coloured solution
• l = path length of light through the solution (cuvette thickness)

Two parts of the law:

1. Lambert's law: Absorbance is directly proportional to the path length of the light through the medium, provided concentration is constant. $A \propto l \quad (\text{when } c \text{ is constant})$

2. Beer's law: Absorbance is directly proportional to the concentration of the coloured substance, provided path length is constant. $A \propto c \quad (\text{when } l \text{ is constant})$

Working principle in a colorimeter:

• A light source (lamp) produces white light, which is made monochromatic by a filter of suitable wavelength.

2. This monochromatic light is passed through a cuvette containing the coloured sample solution.
3. Part of the light is absorbed by the coloured substance and the remaining light is transmitted.
4. A photocell / photodetector measures the transmitted light intensity (Iₜ) and compares it with the incident intensity (I₀) to give absorbance or % transmittance.
5. Since absorbance is proportional to concentration, the unknown concentration is found by comparing with a standard (using a calibration curve or formula):

$c_{\text{unknown}} = \frac{A_{\text{unknown}}}{A_{\text{standard}}} \times c_{\text{standard}}$

Conditions (limitations) for the law to hold:

• Light used must be monochromatic.
• Solution must be dilute (low concentration).
• The coloured substance should not undergo dissociation, association or reaction with the solvent.
• Temperature and pH should remain constant.
• Cuvette must be clean and of uniform path length.

Conclusion: The law of colorimeter (Beer–Lambert law) gives the relation between absorbance, concentration and path length, and forms the basis for quantitative estimation of coloured substances in a sample.

Sample collection technique (1 mark)

• Collect blood by clean venipuncture, preferably from the antecubital vein.
• Use a plain red-top tube or serum separator tube (SST). Do not use EDTA, citrate, oxalate or fluoride tubes, as these can interfere with amylase activity. Heparinised plasma may be acceptable in some methods, but serum is preferred.
• Collect 2–5 mL of blood. Fasting is not mandatory, but avoid alcohol for 24 hours before collection if possible.
• Allow the blood to clot at room temperature for 15–30 minutes. Centrifuge promptly and separate the serum into a clean, labelled tube.
• Label the sample with patient name, date, and exact time of collection, especially in suspected acute pancreatitis.

Precautions during the S. amylase test (1 mark)

• Avoid haemolysed, lipaemic or icteric samples; they may interfere with the enzymatic reaction.
• Do not use EDTA/fluoride/oxalate anticoagulants, as they chelate calcium or inhibit enzyme activity.
• Prevent saliva contamination: saliva is rich in amylase. Do not mouth-pipette; use separate pipettes and keep tubes capped.
• Check expiry date and storage conditions of reagents and calibrator. Equilibrate reagents to room temperature before use.
• Calibrate the analyser and run normal and abnormal quality-control sera before testing the batch.
• Maintain the correct incubation temperature, usually 37°C, and correct wavelength as per the kit method.
• If testing is delayed, separate serum and store at 2–8°C; for longer storage, freeze at −20°C. Avoid repeated freeze–thaw cycles.
• Record drugs that may affect amylase, such as morphine, codeine, diuretics and oral contraceptives.

Interpretation of findings (3 marks)

Important points

• In acute pancreatitis, serum amylase usually rises within 6–12 hours, peaks at 24–48 hours, and returns to normal in 3–5 days.
• A normal amylase does not exclude pancreatitis; correlate with clinical features, lipase, ultrasound and other investigations.
• Isoenzyme study helps to differentiate: P-type (pancreatic) rises in pancreatic disease, while S-type (salivary) rises in salivary gland disease.
• Persistently high serum amylase with low urine amylase suggests macroamylasemia, which is usually a benign condition.

Preparation of Molar and Normal Solutions

1. Molar solution

A molar solution contains one gram-molecular weight (mole) of solute dissolved in one litre of solution. It is denoted by M.

Formula: $M = \frac{\text{weight of solute (g)}}{\text{molecular weight} \times \text{volume of solution (L)}}$

Steps:

1. Calculate the required weight of the solute using the above formula.
2. Weigh the calculated amount of solute accurately on a chemical balance.
3. Transfer it to a clean volumetric flask and dissolve it in a small volume of distilled water.
4. Shake well, then make up the volume exactly to the one-litre mark with distilled water.
5. Stopper the flask, mix thoroughly and label it with the name, molarity and date.

Example: To prepare 1 M sodium chloride (NaCl) solution, the molecular weight of NaCl is 58.5. Weigh 58.5 g of NaCl, dissolve it in distilled water and make the final volume 1 litre. For 0.1 M NaCl, weigh 5.85 g and make up to 1 litre.

2. Normal solution

A normal solution contains one gram-equivalent weight of solute per litre of solution. It is denoted by N.

Formula: $N = \frac{\text{weight of solute (g)}}{\text{equivalent weight} \times \text{volume of solution (L)}}$

where equivalent weight = molecular weight ÷ valency factor.

Steps:

1. Calculate the equivalent weight of the solute.
2. Weigh the required amount and dissolve it in distilled water in a volumetric flask.
3. Make up the volume to the one-litre mark, mix well and label the flask.

Example: To prepare 1 N sodium carbonate (Na₂CO₃) solution, the molecular weight is 106 and the valency factor is 2, so the equivalent weight is 53. Weigh 53 g of Na₂CO₃, dissolve it and make the volume 1 litre. Similarly, 1 N NaOH solution is prepared by dissolving 40 g of NaOH (equivalent weight = 40) in distilled water to make 1 litre.

Note: For dilute acids and bases, the normality equation N₁V₁ = N₂V₂ is used to calculate the volume of the concentrated stock solution required.

VDRL Test (Venereal Disease Research Laboratory Test)

Principle The VDRL test is a non-treponemal flocculation (precipitation) test used for the serological screening of syphilis. It is based on the principle that in syphilis infection, the host tissues release lipoidal material which stimulates the body to produce reagin antibodies (IgM and IgG). These reagin antibodies react not only with the treponemal antigen but also with cardiolipin–lecithin–cholesterol antigen extracted from beef heart. When patient serum containing reagin is mixed with this antigen suspension, the antibodies combine with the antigen particles and form visible floccules (clumps). The cardiolipin acts as the reactive lipid, lecithin enhances the reaction, and cholesterol increases the sensitivity of the test. The test is thus an antigen–antibody precipitation reaction where clumping indicates a positive result.

Procedures

1. Specimen collection: Collect 2–3 ml of venous blood in a clean dry tube without anticoagulant. Allow it to clot, then centrifuge to obtain clear serum. Serum should be tested within 24 hours or stored at 2–8 °C.
2. Inactivation: Heat the serum at 56 °C for 30 minutes in a water bath to inactivate complement and prevent false results.
3. Preparation of antigen: Reconstitute the VDRL antigen (cardiolipin–lecithin–cholesterol) and dilute it with buffered saline as per the manufacturer's instruction. Prepare fresh antigen suspension daily.
4. Slide preparation: Using a wax pencil, draw circles of about 14 mm diameter on a glass slide. Place 0.05 ml of inactivated serum in each circle.
5. Adding antigen: Add one drop (1/60 ml) of freshly prepared antigen suspension to each serum drop using a calibrated needle and syringe.
6. Rotation: Rotate the slide on a mechanical rotator at 180 rpm for 4 minutes (or by hand for 8 minutes).
7. Observation: Examine the slide under a light microscope at 100× magnification for flocculation.
8. Reporting: Report as Reactive, Weakly Reactive or Non-reactive.
9. Quantitative test: If reactive, perform serial two-fold dilution of serum (1:2, 1:4, 1:8 …) and repeat the test to determine the titre, which indicates the antibody level.

Uses

1. Screening of syphilis — used as a routine screening test for syphilis in blood banks, antenatal clinics and preoperative patients.
2. Diagnosis of syphilis — helps in diagnosing primary, secondary and latent syphilis.
3. Monitoring treatment — the fall in titre after treatment indicates cure; a rising titre indicates relapse or reinfection.
4. Screening of blood donors — to prevent transmission of syphilis through blood transfusion.
5. Antenatal screening — to detect syphilis in pregnant women and prevent congenital syphilis.
6. Detection of biological false positives — seen in autoimmune diseases, leprosy, malaria, pregnancy and viral infections.

Note: VDRL is a non-treponemal test, so it may give false-positive results; therefore, a reactive VDRL should be confirmed by a treponemal test such as TPHA or FTA-ABS.

Ten items sterilised in a hot air oven

1. Glass test tubes
2. Glass Petri dishes
3. Glass pipettes
4. Glass beakers and conical flasks
5. All-glass syringes
6. Glass slides and cover slips
7. Metal forceps and scalpels
8. Metal scissors and needles
9. Glass bottles and vials
10. Porcelain mortars, pestles and crucibles

Hot air oven uses dry heat at $160\,^\circ$C for 2 hours (or $170\,^\circ$C for 1 hour). It suits only heat-stable dry items such as glassware, metal instruments, powders, oils and waxes, and not rubber, plastic, cloth or liquid culture media.

Urine analysis by strip (dipstick) method

Principle: The strip carries several reagent pads on a plastic backing. Each pad contains chemicals that react with a specific substance in urine and change colour. The colour change is compared with the colour chart printed on the strip container to get the result.

Procedure:

1. Collect fresh midstream urine in a clean, dry, labelled container. Mix it well and use it within 1–2 hours (older urine gives false results due to bacterial growth and cell lysis).
2. Bring the strip to room temperature and take out only the required number of strips; close the container tightly at once to protect the pads from moisture.
3. Dip the strip completely into the urine so that all pads are wet, then remove it immediately. Do not soak it longer than the time stated on the label (usually 1–2 seconds).
4. Remove excess urine by touching the edge of the strip to the rim of the container or by blotting on absorbent paper. Do not shake or tap the strip, as this mixes reagents between pads.
5. Hold the strip horizontally and read the pads against the colour chart at the correct time for each pad (usually 30–60 seconds; leucocytes about 1–2 minutes). Reading too early or too late gives a wrong result.
6. Record the findings in the report form: colour, clarity, pH, specific gravity, protein, glucose, ketones, blood, bilirubin, urobilinogen, nitrite and leucocytes.
7. Compare the result with the reference range and report as negative, trace, 1+, 2+, 3+ or 4+ as given on the chart. If any pad is positive, confirm by a chemical or microscopic method.

Precautions: Use strips before the expiry date, keep them in a cool dry place in the closed container, do not touch the pads with fingers, and always use a fresh, well-mixed sample.

Benefits of the strip method:

1. Simple and quick — the whole test is finished in about 1–2 minutes and needs no special skill or laboratory set-up.
2. Screening of many tests at once — one strip tests up to 10 parameters (pH, protein, glucose, ketones, blood, bilirubin, etc.) at the same time.
3. Cheap and portable — low cost per test, small size, so it can be used in health posts, camps and field conditions.
4. Safe and hygienic — no corrosive liquid reagents, no heating or centrifugation, so there is little risk to the worker.
5. Useful for early detection and follow-up — helps in screening diabetes (glucose), kidney disease (protein, blood), urinary infection (nitrite, leucocytes) and liver disease (bilirubin), and in monitoring patients repeatedly.

Parts of a compound microscope and their uses in handling

A compound microscope magnifies small objects using two lens systems, the objective and the eyepiece.

A. Mechanical parts

• (a) Base: the heavy foot that keeps the instrument steady; support it with one hand when carrying.
• (b) Arm (limb): the curved part joining base and head; hold it as the handle when carrying. It carries the stage and the focusing knobs.
• (c) Stage: the flat platform with a central hole on which the slide is placed; light passes through the hole to the specimen.
• (d) Stage clips (or mechanical stage): hold the slide in place and let it be moved smoothly during scanning.
• (e) Body tube: the tube holding the eyepiece above and the objectives below at the correct distance.
• (f) Draw tube: a sliding tube at the top of the body tube used to adjust the tube length (distance between eyepiece and objective); it is not used for focusing.
• (g) Coarse adjustment knob: the large knob that moves the tube or stage quickly for first focusing, with the low-power objective only. Turn it while watching from the side to avoid crashing the objective onto the slide.
• (h) Fine adjustment knob: the small knob for final, sharp focusing; it is the only focusing knob used with the high-power and oil-immersion objectives.
• (i) Revolving nosepiece: the rotating disc carrying the objectives; turn it by its rim, not by the lenses, to change magnification.

B. Optical and illuminating parts

• (j) Objective lenses: usually $4\times$ or $10\times$ (low power), $40\times$ (high dry) and $100\times$ (oil immersion). They form the first magnified image. Clean only with lens paper; remove oil after using $100\times$.
• (k) Eyepiece (ocular): usually $10\times$; it further magnifies the image. Total magnification = objective power $\times$ eyepiece power.
• (l) Mirror or built-in lamp: directs light up through the specimen. Use the plane side in bright light and the concave side in dim light.
• (m) Condenser: focuses light onto the specimen; raise it for high power and lower it for low power.
• (n) Iris diaphragm: controls the amount of light and the contrast.

C. Rules for handling

1. Carry with one hand on the arm and the other under the base.
2. Place on a firm, level table away from its edge.
3. Always begin with the low-power objective, focus with the coarse knob, then finish with the fine knob.
4. Never touch lenses with fingers; use only lens paper, and wipe oil off the $100\times$ objective after use.
5. After use, return to the low-power objective, lower the stage or tube, remove the slide, and cover the microscope.

Interpretation of results and sources of error in blood urea estimation

Urea is the main end product of protein metabolism, formed in the liver and excreted by the kidneys. Common methods are the urease-Berthelot (indophenol), urease-GLDH (enzymatic UV) and diacetyl monoxime methods.

A. Interpretation of results

Normal serum urea in adults is about 15-45 mg/dL (blood urea nitrogen, BUN, about 7-20 mg/dL). Urea = BUN $\times$ 2.14.

Raised urea (azotaemia)

Low urea: severe liver disease, low-protein diet or malnutrition, pregnancy, over-hydration, SIADH.

The BUN : creatinine ratio helps localise the cause. A ratio above 20 : 1 suggests pre-renal azotaemia, about 10-20 : 1 suggests intrinsic renal disease, and below 10 : 1 suggests liver disease or low protein intake. Urea is influenced by diet and hydration, so it should always be read together with creatinine and clinical findings.

B. Sources of error

Pre-analytical

• Use of ammonium-containing anticoagulants (ammonium oxalate or ammonium heparin) or ammonia contamination of glassware, which falsely raises urease-Berthelot results.
• Haemolysed, lipaemic or icteric samples.
• Delay in separating serum, or storage at room temperature: bacterial urease breaks down urea and so lowers the measured value (and raises ammonia).
• Wrong sample labelling; sampling after a heavy protein meal or when dehydrated.

Analytical

• Unchecked calibration or an expired or wrongly prepared standard.
• Pipetting errors and dirty glassware or cuvettes.
• Deteriorated or contaminated reagents (urease, phenol, hypochlorite), or wrong buffer pH.
• Wrong incubation time or temperature; reading after the colour has faded.
• Interference from ammonia in the sample or reagents, and from bilirubin and haemoglobin in some methods.
• Carry-over between samples in automated analysers.

Post-analytical

• Calculation errors, wrong unit or wrong BUN-to-urea conversion.
• Transcription errors and use of the wrong reference range.

C. Precautions

Use a suitable anticoagulant (heparin without ammonium salts, or serum), separate and analyse promptly or store at 2-8 $^\circ$C, avoid haemolysis, calibrate regularly, run control sera with each batch, use fresh reagents, and cross-check abnormal results before reporting.