Direct Coombs's test (Direct Antiglobulin Test, DAT)

Principle: The test detects antibodies or complement components already coated on the surface of the patient's red blood cells in vivo. Anti-human globulin (Coombs's) reagent, which is an antibody against human IgG and/or complement (C3d), bridges these sensitized red cells and causes visible agglutination. It is called "direct" because sensitization has already occurred inside the body and no serum is added in vitro.

Reagents and materials required:

• Anti-human globulin (Coombs's) reagent — polyspecific (anti-IgG + anti-C3d) or monospecific
• Normal saline (0.9% NaCl), pH 6.8–7.2
• Test tubes (10 × 75 mm), Pasteur pipettes, centrifuge, water bath (37 °C), glass slide, light source/mirror
• Patient's blood sample collected in EDTA (or plain tube) — preferably within 24 hours

Procedure:

1. Collect 3–5 mL of venous blood in EDTA. Centrifuge at 1500–2000 rpm for 5 minutes to obtain packed red cells.
2. Wash the red cells 3–4 times with a large volume of normal saline. After each wash, centrifuge and discard the supernatant completely. This removes plasma proteins (especially IgG) that would otherwise neutralize the anti-human globulin reagent.
3. Prepare a 2–5% red cell suspension in normal saline from the washed packed cells.
4. Label a clean test tube and place one drop of the 2–5% red cell suspension into it.
5. Add one drop of anti-human globulin (Coombs's) reagent to the tube.
6. Mix gently and centrifuge at 1000–1500 rpm for 1 minute (or 3000 rpm for 15–20 seconds).
7. Gently resuspend the cell button and examine for agglutination macroscopically against a well-lit white background; confirm microscopically if weak.
8. Controls:
   • Positive control: known IgG-sensitized red cells — must show agglutination.
   • Negative control: unsensitized (Coombs-negative) red cells — must show no agglutination.
   • Saline control: red cells + saline only — rules out spontaneous agglutination.
   • If negative, add one drop of known IgG-sensitized cells to confirm the reagent was not neutralized (validity check).

Interpretation:

• Positive (agglutination): red cells are coated with IgG and/or complement in vivo — seen in autoimmune haemolytic anaemia, haemolytic disease of the newborn (Rh incompatibility), drug-induced haemolysis, and delayed haemolytic transfusion reactions.
• Negative (no agglutination): no significant in vivo sensitization; if clinically suspected, repeat with monospecific reagents and a fresh sample.

Precautions:

• Wash red cells thoroughly; incomplete washing gives false-negative results.
• Use fresh reagents and check expiry; use saline at correct pH.
• Read results immediately after centrifugation; delayed reading may give false-positive results due to drying.
• Avoid contamination and use only clean glassware.

Composition and Function of Plasma [5 marks]

Plasma is the pale yellow, slightly alkaline (pH 7.4) liquid portion of blood, forming about 55% of total blood volume. It is obtained when blood is centrifuged after adding an anticoagulant. It is a complex mixture of water and dissolved substances.

Composition of Plasma

Functions of Plasma

1. Transport: Carries nutrients (glucose, amino acids, lipids), hormones, enzymes, vitamins, and waste products (urea, uric acid) to and from tissues; also transports O₂ and CO₂ in dissolved form.
2. Regulation of osmotic pressure: Plasma proteins, mainly albumin, maintain osmotic (oncotic) pressure and thus keep water balance between blood and tissues.
3. Regulation of pH: Buffer systems of plasma (bicarbonate, phosphate, and plasma proteins) keep blood pH constant at about 7.4.
4. Defence: Globulins (antibodies) provide immunity against infection; plasma also carries white blood cells and clotting factors.
5. Coagulation: Fibrinogen is essential for blood clotting, preventing blood loss from injured vessels.
6. Temperature regulation: Water in plasma absorbs and distributes heat, helping to maintain normal body temperature.
7. Maintenance of blood volume and pressure: Plasma provides the fluid medium that keeps blood volume and hence blood pressure normal.

Calculation of RBC Indices

RBC indices are calculated from three directly measured values — haemoglobin (Hb), packed cell volume (PCV/haematocrit) and RBC count — either manually or automatically by a haematology analyser. The three main indices are MCV, MCH and MCHC; RDW is also reported.

1. Mean Corpuscular Volume (MCV)

Average volume of a single red cell, expressed in femtolitres (fL).

$\text{MCV} = \frac{\text{PCV (\%)} \times 10}{\text{RBC count (million/µL)}} \text{ fL}$

2. Mean Corpuscular Haemoglobin (MCH)

Average weight of haemoglobin in one red cell, expressed in picograms (pg).

$\text{MCH} = \frac{\text{Hb (g/dL)} \times 10}{\text{RBC count (million/µL)}} \text{ pg}$

3. Mean Corpuscular Haemoglobin Concentration (MCHC)

Concentration of haemoglobin in a given volume of packed red cells, expressed in g/dL or %.

$\text{MCHC} = \frac{\text{Hb (g/dL)} \times 100}{\text{PCV (\%)}} \text{ g/dL}$

4. Red Cell Distribution Width (RDW)

Coefficient of variation of red cell size, reported as a percentage.

$\text{RDW} = \frac{\text{SD of MCV}}{\text{Mean MCV}} \times 100$

Worked Example

Given: Hb = 12 g/dL, PCV = 40 %, RBC count = 4.5 million/µL

• MCV = (40 × 10) / 4.5 = 88.9 fL
• MCH = (12 × 10) / 4.5 = 26.7 pg
• MCHC = (12 × 100) / 40 = 30 g/dL

Normal Reference Values

Interpretation

• MCV – low in microcytic anaemia (iron deficiency, thalassaemia), high in macrocytic anaemia (B₁₂/folate deficiency).
• MCH – low in hypochromic anaemia; roughly parallels MCV.
• MCHC – low in iron deficiency and thalassaemia; high in spherocytosis and sickle cell disease.
• RDW – raised when there is anisocytosis; helps distinguish iron deficiency (high RDW) from thalassaemia trait (normal RDW).

Derived Index (for differentiation)

$\text{Mentzer index} = \frac{\text{MCV}}{\text{RBC count}}$ Value < 13 suggests thalassaemia trait; > 13 suggests iron deficiency anaemia.

RPR test vs VDRL test (both are non-treponemal flocculation tests for syphilis screening)

Key point: Both detect reagin (antibody) and become negative after successful treatment; a reactive result in either must be confirmed by a treponemal test (TPHA/FTA-ABS).

Gram's stain — principle and procedure

Principle (2 marks)

Gram's stain differentiates bacteria into Gram-positive and Gram-negative groups on the basis of the structure of their cell wall.

• Gram-positive bacteria have a thick peptidoglycan layer (about 20–80 nm) with little lipid. The primary stain crystal violet forms a large crystal violet–iodine (CV-I) complex inside the cell. During decolourisation with alcohol/acetone, the thick peptidoglycan layer dehydrates, shrinks and closes its pores, so the CV-I complex is trapped and cannot be washed out. The cell therefore retains the violet colour even after the counterstain.
• Gram-negative bacteria have a thin peptidoglycan layer (about 2–3 nm) covered by a lipid-rich outer membrane. The alcohol dissolves the outer membrane lipids and damages the thin peptidoglycan, so the CV-I complex leaches out and the cell becomes colourless. It then takes up the pink/red colour of the counterstain (safranin or dilute carbol fuchsin).

Thus the difference in cell wall composition (peptidoglycan thickness and lipid content) is the basis of the reaction. Gram-positive = purple/violet; Gram-negative = pink/red.

Procedure (3 marks)

1. Preparation of smear: Place a loopful of saline/water on a clean grease-free slide. Emulsify a small portion of a young (18–24 hour) culture or specimen and spread it into a thin film. Air-dry and heat-fix by passing the slide over a flame 2–3 times (do not overheat).
2. Primary stain — crystal violet: Flood the smear with crystal violet (gentian violet) and leave for 1 minute. Wash gently with water.
3. Mordant — Gram's iodine: Flood with Gram's iodine and leave for 1 minute. The iodine combines with crystal violet to form the CV-I complex. Wash with water.
4. Decolourisation: Apply 95% ethyl alcohol (or acetone–alcohol) drop by drop until the run-off becomes colourless (about 5–15 seconds). Wash immediately with water. This is the critical step — over-decolourisation gives false Gram-negative results.
5. Counterstain: Flood with safranin (or dilute carbol fuchsin) for 30 seconds to 1 minute. Wash with water.
6. Drying and examination: Blot dry (do not rub) or air-dry, and examine under the oil-immersion objective (100×).

Result: Gram-positive organisms appear deep purple/violet; Gram-negative organisms appear pink/red.

Stool Examination by Wet Mount Preparation

Principle: A wet mount (direct saline and iodine preparation) is a simple, rapid microscopic method used to detect ova, cysts, larvae, trophozoites and motile bacteria in stool. Saline reveals motility and morphology; iodine stains glycogen and nuclei for clearer identification of cysts.

Materials Required

• Fresh stool specimen (preferably within 30 minutes of passage)
• Clean glass slides and coverslips
• Normal saline (0.85% NaCl)
• Lugol's iodine (1%)
• Applicator sticks / wooden sticks
• Gloves, marker, microscope

Procedure

1. Label a clean glass slide with the patient's name and date.
2. Saline mount: Place one drop of normal saline on the left half of the slide. With an applicator stick, take a small portion of stool (about the size of a match head) from the mucus/blood-tinged area and mix it in the saline to make a thin, uniform suspension.
3. Iodine mount: Place one drop of Lugol's iodine on the right half of the slide. Emulsify a similar portion of stool in it.
4. Coverslip: Place a coverslip gently over each preparation, avoiding air bubbles and overflow.
5. Microscopic examination: First examine under low power (10×) to scan the entire field for ova, cysts and larvae. Then confirm under high power (40×) for detailed morphology and motility.
6. Reporting: Report the findings as "ova/cyst seen" or "no ova/cyst seen" with the type identified.

Precautions

• Use a fresh specimen; delay causes trophozoites to disintegrate and cysts to distort.
• Do not make the preparation too thick or too thin.
• Avoid contamination with urine or water.
• Examine within 30 minutes for motile trophozoites.
• Wear gloves and disinfect the working area properly.

(a) Methods of hemoglobin estimation — 2 marks

1. Colorimetric / photometric methods
   • Cyanmethemoglobin (HiCN) method — reference method
   • Oxyhemoglobin method
   • Alkaline hematin method
   • Acid hematin method (Sahli's method)
2. Gasometric method — Van Slyke apparatus (measures oxygen combining capacity)
3. Chemical method — Iron content estimation (Drabkin's iron method)
4. Specific gravity method — Copper sulphate falling drop method
5. Direct reading methods — Sahli's hemoglobinometer, Dare's hemoglobinometer, Tallqvist's scale
6. Automated methods — Spectrophotometry in automated cell counters (e.g., Sysmex), which use the sodium lauryl sulphate (SLS) or cyanmethemoglobin principle

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(b) Cyanmethemoglobin (HiCN) method — principle — 2 marks

Principle:

• Blood is diluted in Drabkin's solution, which contains potassium ferricyanide K₃Fe(CN)₆ and potassium cyanide (KCN).
• Ferricyanide oxidises haemoglobin (Hb) to methaemoglobin (Hi): $\text{Hb} + \text{K}_3\text{Fe(CN)}_6 \rightarrow \text{Hi} + \text{K}_4\text{Fe(CN)}_6$
• Methaemoglobin then combines with cyanide to form cyanmethemoglobin (HiCN), a stable coloured compound: $\text{Hi} + \text{KCN} \rightarrow \text{HiCN}$
• The absorbance of HiCN is measured at 540 nm in a spectrophotometer/colorimeter.
• Colour intensity is directly proportional to Hb concentration (Beer–Lambert law).

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(c) Procedure of cyanmethemoglobin method — 3 marks

1. Reagents:
   • Drabkin's solution: potassium ferricyanide 200 mg, potassium cyanide 50 mg, potassium dihydrogen phosphate 140 mg, distilled water to 1000 mL (pH 7.0–7.4).
   • HiCN standard (known Hb concentration, e.g., 15 g/dL).
2. Sample: Well-mixed venous blood (EDTA anticoagulant).
3. Dilution: Take 20 µL of blood into 5 mL of Drabkin's solution (dilution 1:251) in a clean test tube. Mix gently and let stand at room temperature for 5–10 minutes for full colour development.
4. Blank: Take 5 mL Drabkin's solution in another tube (no blood).
5. Standard: Treat the HiCN standard in the same way as the sample.
6. Measurement: Set the colorimeter/spectrophotometer to 540 nm, zero the instrument with the blank. Read the absorbance (OD) of the standard and the test.
7. Calculation: $\text{Hb (g/dL)} = \frac{\text{OD of test}}{\text{OD of standard}} \times \text{Concentration of standard (g/dL)}$ Or using the millimolar extinction coefficient (per haem, $\varepsilon_{540} = 11.0\ \text{L mmol}^{-1}\text{cm}^{-1}$, light path 1 cm): $\text{Hb (mmol/L, as haem)} = \frac{A_{540} \times 251}{11.0 \times 1}$ Converting with the haem-equivalent weight of Hb (64,500 / 4 = 16,125 g/mol, i.e. 1 g/dL = 0.62 mmol/L haem): $\text{Hb (g/dL)} = \frac{A_{540} \times 251}{11.0 \times 0.62} \approx A_{540} \times 36.8$ Example: $A_{540} = 0.40$ gives Hb of about 14.7 g/dL.

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(d) Drawbacks of cyanmethemoglobin method — 3 marks

1. Toxicity of reagents: Drabkin's solution contains potassium cyanide, which is highly poisonous; it can release HCN gas on contact with acids. Strict precautions and proper disposal are needed.
2. Interference by lipaemia and turbidity: Lipaemic or icteric plasma and high WBC/platelet counts cause turbidity, falsely raising absorbance. A turbidity blank (sample + Drabkin's solution without ferricyanide) is required.
3. Interference by other Hb derivatives:
   • Carboxyhaemoglobin (HbCO) is not converted to HiCN and gives falsely low values.
   • Sulfhaemoglobin (SHb) is not measured and gives falsely low values.
   • Methaemoglobin is measured but with a slightly different extinction coefficient, causing minor error.
4. Not suitable for field use: Requires a spectrophotometer/colorimeter, accurate pipettes and a stable standard; hence unsuitable for bedside or rural field conditions.
5. Standard instability: HiCN standards are light-sensitive and deteriorate with time; improper storage gives erroneous results.
6. Longer time required: Needs 5–10 minutes for complete colour development, unlike Sahli's method which is quicker.
7. Disposal problem: Cyanide-containing waste is an environmental hazard and must be treated (e.g., with sodium hypochlorite) before disposal.

Methods of Sterilization

1. Physical methods

• Heat
  • Dry heat: Red heat (inoculating loops), flaming, hot air oven (160–180°C for 1–2 hours), incineration.
  • Moist heat: Boiling (100°C, 10–30 min), autoclave (steam under pressure, 121°C at 15 psi for 15–20 min), pasteurization, tyndallization (intermittent steaming).
• Radiation
  • Non-ionizing: Ultraviolet light (surface and air disinfection).
  • Ionizing: Gamma rays, X-rays, electron beams (cold sterilization of disposables).
• Filtration: Membrane filtration (0.22 µm), sintered glass, Seitz filter — for heat-labile fluids, sera, antibiotics, vaccines.

2. Chemical methods

• Liquids: Alcohol (70% ethanol, isopropanol), formaldehyde, glutaraldehyde (2%), hydrogen peroxide, phenol, chlorine compounds, iodophors, quaternary ammonium compounds.
• Gases: Ethylene oxide, formaldehyde vapour, hydrogen peroxide vapour, plasma sterilization — for heat-sensitive instruments.

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Autoclave

Principle: Moist heat under pressure. Steam under pressure raises the boiling point of water, so saturated steam at 121°C (15 psi / 1.05 kg/cm²) kills all microorganisms including spores within 15–20 minutes by coagulation of proteins.

Structure: A strong double-walled cylindrical vessel of stainless steel with an outer jacket and inner chamber. Fitted with a pressure gauge, safety valve, steam inlet and outlet valves, thermometer, and a lid with locking clamps. Modern autoclaves are horizontal, front-loading, and may be fully automatic.

Types:

• Simple/bench autoclave — manually operated, gravity displacement.
• Horizontal/vertical autoclave — for bulk loads.
• Vacuum (pre-vacuum) autoclave — air removed by vacuum pump, faster steam penetration.
• Flash autoclave — 134°C for 3 minutes for urgent items.

Working (gravity displacement):

1. Water is filled in the chamber (or steam is admitted from a boiler).
2. Lid is closed and clamped; outlet valve is kept open.
3. Heating drives out air; when steam escapes freely from the outlet, the valve is closed.
4. Pressure rises to 15 psi, temperature to 121°C; timing starts.
5. Load is held for 15–20 min (varies with load).
6. Heat is stopped, pressure allowed to fall to zero, outlet opened, lid opened slowly.

Uses: Sterilization of dressings, linen, surgical instruments, glassware, culture media, rubber gloves, and infectious waste.

Precautions:

• Load must not be packed too tightly; air must be completely removed.
• Correct pressure–temperature–time relation must be maintained.
• Lid must not be opened before pressure falls to zero.
• Glassware should be loosely capped; liquids in vented containers.

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Quality Control of Autoclave

1. Physical (mechanical) monitoring

• Record pressure gauge, thermometer, and time for every cycle.
• Check for steam leaks, faulty valves, and worn gaskets.
• Compare the reading with a calibrated reference thermometer.

2. Chemical monitoring

• Autoclave tape — changes colour (e.g., white to black stripes) when exposed to steam.
• Bowie–Dick test — a test pack placed in the centre of the load; uniform colour change confirms steam penetration and proper air removal. Done daily/with each load.
• Chemical indicator strips / Browne's tubes — colour change confirms exposure to required temperature and steam.

3. Biological monitoring (the most reliable)

• Geobacillus stearothermophilus (formerly Bacillus stearothermophilus) spores are used as the biological indicator.
• Spore strips or ampoules are placed in the load, incubated at 55–60°C for 7 days along with a control.
• No growth in the test and growth in the control = satisfactory sterilization.
• Growth in the test = sterilization failure; the load must be reprocessed and the autoclave checked.
• Should be done at least weekly and after every major repair.

4. Other checks

• Regular cleaning and descaling of the chamber.
• Annual calibration of gauges and safety valves.
• Maintenance of a logbook of every cycle and test result.

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Note: Proper loading, complete air removal, and regular biological monitoring are essential, because air pockets prevent steam contact and give false assurance of sterility.

CSF Examination in the Laboratory

CSF (Cerebrospinal Fluid) examination is done in three main stages: collection and physical examination, chemical examination, and microscopic examination.

1. Collection and Handling

• CSF is collected by lumbar puncture (usually between L3–L4 or L4–L5) by a physician.
• Collected in three sterile numbered tubes (2–3 mL each):
  • Tube 1 — for chemical and serological tests (least likely to contain blood from the puncture).
  • Tube 2 — for microbiological culture and staining.
  • Tube 3 — for cell count (least contamination from puncture trauma).
• Specimen must be examined immediately (within 1 hour) because cells lyse quickly; if delayed, refrigerate at 2–8 °C (never freeze).

2. Physical Examination

• Appearance: Note whether clear, cloudy/turbid (↑ cells, protein), bloody (trauma/subarachnoid haemorrhage), or xanthochromic (yellow — old bleed, jaundice).
• Volume, colour, and viscosity are also noted.
• Clot formation: Normal CSF does not clot; a fibrin web/web clot may form in tubercular meningitis (↑ fibrinogen).

3. Chemical Examination

• Protein: Measured by Pandy's test (qualitative) or quantitative methods (Biuret / turbidimetry). Normal: 15–45 mg/dL.
• Glucose: Measured by GOD-POD method. Normal: 45–80 mg/dL (about 60% of blood glucose). Decreased in bacterial and tubercular meningitis.
• Chloride: Estimated by titration (Schales and Schales). Normal: 700–750 mg/dL. Decreased in tubercular meningitis.
• Lactate and LDH may also be estimated when indicated.

4. Microscopic Examination

• Cell count: Done in a Fuchs–Rosenthal counting chamber (or Neubauer chamber) using undiluted CSF. Normal: 0–5 lymphocytes/µL in adults.
• Differential count: After cytocentrifugation or staining with Leishman / Giemsa stain, count lymphocytes, neutrophils, and other cells.
• Microscopy for organisms:
  • Gram stain for bacteria.
  • Ziehl–Neelsen stain for Mycobacterium tuberculosis.
  • India ink preparation for Cryptococcus neoformans.
• Culture: CSF is inoculated on blood agar, chocolate agar, and MacConkey agar, and in Robertson's cooked meat medium for anaerobes.

5. Interpretation (Typical Findings)

Conclusion: A properly collected, promptly processed CSF sample with physical, chemical, and microscopic examination provides the key diagnostic information for meningitis and other CNS disorders.

Beer's and Lambert's law (also called Beer–Lambert law) 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 through which light passes.

$A = \varepsilon \cdot c \cdot l$

Where:

• A = absorbance (optical density, OD)
• ε = molar absorptivity (a constant for a given substance at a given wavelength)
• c = concentration of the substance
• l = path length of light through the solution (cuvette width, usually 1 cm)

Combined statement:

• Lambert's law: Absorbance is directly proportional to the path length (l) of the solution.
• Beer's law: Absorbance is directly proportional to the concentration (c) of the coloured substance.

Application in clinical laboratory:

1. Colorimetry / Spectrophotometry: Used to estimate concentration of substances in body fluids (blood glucose, serum bilirubin, urea, creatinine, cholesterol, haemoglobin, etc.) by measuring absorbance of coloured solutions.

2. Quantitative estimation: Since A ∝ c (when l and ε are constant), the unknown concentration is calculated by comparing with a standard of known concentration:

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

3. Calibration curve: A standard curve of absorbance versus concentration is plotted; the absorbance of the test sample is read against it to find its concentration.

4. Quality control: Used to check linearity, reagent stability and instrument performance in the laboratory.

5. Choice of wavelength: The wavelength of maximum absorbance (λmax) is selected so that small changes in concentration produce measurable changes in absorbance, giving accurate results.

Functions of Blood (5 marks)

Blood performs the following major functions:

1. Transportation

   • Carries oxygen from the lungs to the tissues (by haemoglobin in RBCs) and carbon dioxide from tissues back to the lungs.
   • Transports nutrients (glucose, amino acids, fatty acids, vitamins, minerals) absorbed from the digestive tract to all body cells.
   • Carries hormones from endocrine glands to their target organs.
   • Transports metabolic wastes (urea, uric acid, creatinine) to the kidneys, and heat to the skin for elimination.

2. Regulation

   • Regulates body temperature by distributing heat and by vasoconstriction/vasodilation of skin vessels.
   • Maintains acid–base balance (pH about 7.4) through buffers such as bicarbonate and plasma proteins.
   • Maintains water and electrolyte balance between blood and tissues.

3. Protection (Defence)

   • White blood cells (neutrophils, lymphocytes, monocytes) phagocytose and destroy invading microorganisms.
   • Lymphocytes produce antibodies and provide immunity against infections.
   • Blood clotting (haemostasis) by platelets and clotting factors prevents blood loss from injured vessels.

4. Maintenance of homeostasis

   • Keeps the internal environment constant by carrying substances between organs and by feedback regulation.

5. Other functions

   • Carries drugs, antibiotics and other therapeutic agents to the tissues.
   • Serves as a reservoir of water, proteins and electrolytes.
   • Helps in the distribution of hormones and enzymes and in defence against toxins.

Structure and Functions of the Gastrointestinal Tract (GI Tract)

Introduction The gastrointestinal tract (GI tract) is a continuous muscular tube, about 9 metres long in an adult, extending from the mouth to the anus. It is also called the alimentary canal. Its main role is to digest food, absorb nutrients, and eliminate waste.

Structure The GI tract consists of the following parts:

1. Mouth (Buccal cavity) – Contains teeth, tongue, and salivary glands. Teeth masticate food; saliva (containing salivary amylase) moistens it and begins starch digestion. The tongue helps in mixing and swallowing.
2. Pharynx and Oesophagus – The pharynx is the common passage for food and air. The oesophagus is a muscular tube about 25 cm long that carries food to the stomach by peristalsis.
3. Stomach – A J-shaped muscular bag with three regions: fundus, body, and pylorus. Its wall has gastric glands secreting HCl, pepsinogen, and mucus. It stores food, churns it, and starts protein digestion.
4. Small intestine – About 6–7 m long, divided into duodenum, jejunum, and ileum. Its inner surface has villi and microvilli that greatly increase the surface area for absorption. The duodenum receives bile from the liver and pancreatic juice from the pancreas.
5. Large intestine – About 1.5 m long, divided into caecum, colon, and rectum. It absorbs water and electrolytes and forms faeces. The rectum stores faeces, which are expelled through the anus.

Histology (wall layers) From outside inward, the GI wall has four layers: serosa, muscularis externa (longitudinal and circular muscles), submucosa, and mucosa. The mucosa contains glands and villi.

Functions

1. Ingestion – Taking food into the mouth.
2. Digestion – Mechanical (chewing, churning) and chemical (enzymes) breakdown of food into simple absorbable forms.
3. Absorption – Passage of digested nutrients, water, and minerals into blood or lymph, mainly in the small intestine.
4. Assimilation – Utilization of absorbed nutrients by body cells.
5. Egestion/Defaecation – Removal of undigested waste as faeces through the anus.
6. Other functions – Secretion of digestive juices, maintenance of water and electrolyte balance, and protection by mucus and gastric acid against microbes.

Conclusion Thus, the GI tract is a well-organized muscular tube whose different parts work together to digest food, absorb nutrients, and expel waste, thereby maintaining nutrition and health of the body.

Features of Different Types of Microscopes

Microscopes are broadly divided into light (optical) microscopes and electron microscopes, based on the source of illumination used to view the specimen.

1. Light (Optical) Microscope

Uses visible light (wavelength ~400–700 nm) and a system of glass lenses to magnify the specimen. Its main types are:

a) Simple (Simple) Microscope

• Consists of a single convex lens (or a pair of lenses acting as one).
• Magnification is low, usually 5× to 20×.
• Simple, cheap and portable; used for preliminary/dissecting work and by students.
• Resolution is poor, so fine cell details cannot be seen.

b) Compound Microscope

• Uses two lens systems — objective and eyepiece — giving higher magnification (40× to 1000×).
• Specimen must be thin and transparent; viewed as a thin section in transmitted light.
• Has a mechanical stage, coarse and fine adjustment knobs, condenser and iris diaphragm.
• Resolution is about 0.2 µm; used to study cells, tissues, bacteria and blood films.
• Stains (e.g., Gram stain, Leishman stain) are needed to increase contrast.

c) Dark-field Microscope

• The specimen is illuminated from the side so that the background remains dark and the object appears bright against a dark background.
• Useful for viewing living, unstained, transparent organisms such as Treponema pallidum (syphilis spirochaete) and leptospira.

d) Phase-contrast Microscope

• Converts differences in refractive index of cell parts into differences in light intensity (contrast).
• Allows observation of living, unstained cells and their internal structures (nucleus, vacuoles, organelles).
• Widely used in tissue-culture and cytology laboratories.

e) Fluorescence Microscope

• Specimen is stained with a fluorescent dye (e.g., auramine, FITC) and viewed under ultraviolet light.
• The object appears glowing against a dark background.
• Used in immunofluorescence (e.g., diagnosis of rabies, autoimmune diseases) and for detecting Mycobacterium tuberculosis by auramine staining.

f) Phase/Interference and Polarising Microscope

• Polarising microscope uses polarised light to study birefringent materials such as crystals, amyloid, collagen and minerals.

2. Electron Microscope

Uses a beam of electrons (wavelength ~0.005 nm) instead of light, so resolution is far greater (0.1–0.5 nm).

a) Transmission Electron Microscope (TEM)

• Electrons pass through an ultra-thin specimen; image formed on a fluorescent screen or photographic plate.
• Very high magnification (up to 1,000,000×) and resolution; shows internal ultrastructure (organelles, viruses).
• Specimen must be fixed, dehydrated, embedded and sectioned; living cells cannot be seen.

b) Scanning Electron Microscope (SEM)

• A beam of electrons scans the surface of the specimen; secondary electrons are collected to form a three-dimensional image.
• Magnification 10× to 100,000×; shows surface topography of cells, insects, pollen, etc.
• Specimen is coated with a thin film of gold or carbon.

Summary Table

Conclusion: The choice of microscope depends on the purpose — light microscopes for routine and living specimens, and electron microscopes for the finest structural details.

Decalcification is the process of removing calcium salts (mainly calcium carbonate and calcium phosphate) from calcified tissues such as bone, teeth, and calcified cartilage so that they become soft enough to be cut into thin sections by the microtome without damaging the knife or tearing the tissue. It is done after fixation and before dehydration, embedding, and sectioning in histopathology.

Methods of decalcification:

1. Acid decalcification (chemical method)

• Strong acids:
  • Nitric acid (5–10% aqueous or 10% formal nitric acid): Rapid action (1–3 days), good for bone marrow and routine bone specimens. Disadvantages: causes swelling of tissue, destroys nuclear staining, and forms nitrous acid fumes that damage tissue and equipment.
  • Hydrochloric acid (5–10%): Very rapid but harsh; causes tissue swelling and poor staining; rarely used alone.
  • Hydrochloric acid–formalin mixture: Combines fixation and decalcification; fast but can distort tissue.
• Weak acids:
  • Formic acid (5–10% aqueous or formic acid–formalin): Most widely used; slower (3–7 days) but gentle, preserves tissue morphology and staining quality well; suitable for routine bone and small specimens.
  • Acetic acid (5%): Very weak and slow; used only for small, mildly calcified tissues; may cause tissue swelling.
  • Trichloroacetic acid (5%): Weak acid, preserves morphology but slow and expensive; used for small specimens.

2. Chelating method (EDTA)

• EDTA (ethylenediaminetetraacetic acid, 10% neutral, pH 7.0–7.4): Chelating agent; very slow (weeks) but the most gentle method — preserves cellular detail, enzymes, and antigens; method of choice for immunohistochemistry, electron microscopy, and research tissues. EDTA binds calcium ions and removes them without acid damage. Best for delicate tissues and special stains.

3. Ion-exchange resin method — the tissue is placed in a decalcifying fluid (e.g., formic acid) along with an ion-exchange resin (e.g., Amberlite IR-120). The resin continuously removes released calcium ions, keeping the solution unsaturated, so decalcification is faster and gentler than acid alone. Good for small bone specimens.

4. Electrolytic (electrophoretic) decalcification — the tissue is placed in an acid decalcifying bath through which a direct electric current is passed; calcium ions migrate to the cathode and are removed. It is the fastest method (hours to a few days) but needs careful control of current, heat, and pH; risk of tissue damage if not monitored.

5. Ultrasonic method — ultrasonic waves agitate the decalcifying fluid and speed up removal of calcium; often combined with acid or EDTA; useful for small specimens.

6. Microwave method — microwave irradiation accelerates acid decalcification; rapid but requires strict temperature control to avoid tissue damage.

Factors affecting decalcification: type and concentration of decalcifying agent, temperature (higher = faster but more damage), agitation, volume of fluid (should be 20–30 times tissue volume), and size/thickness of specimen (2–4 mm thick is ideal).

Assessment of endpoint: chemical test (calcium oxalate/ammonium oxalate test), X-ray examination, physical test (needle/knife cutting), or weight loss method. Once decalcification is complete, tissue is washed in running water to remove acid before processing.

Liver Function Test (LFT)

Liver Function Test (LFT) is a group of biochemical blood tests performed to assess the functional status of the liver, detect liver injury, and help diagnose and monitor liver diseases such as hepatitis, cirrhosis, obstructive jaundice, and drug-induced liver damage.

LFTs are broadly grouped as:

1. Tests of hepatocellular injury (transaminases): ALT (GPT) and AST (GOT).
2. Tests of cholestasis / biliary obstruction: Alkaline phosphatase (ALP), GGT, total and direct bilirubin.
3. Tests of synthetic function: Total protein, albumin, A/G ratio, prothrombin time (PT/INR).
4. Tests of metabolic/detoxifying function: Ammonia, bilirubin metabolism tests.

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Estimation of GPT (ALT) by Kinetic Method

Principle

GPT (Glutamate Pyruvate Transaminase), also called ALT (Alanine Aminotransferase), catalyses the reversible transfer of an amino group from L-alanine to α-ketoglutarate, forming pyruvate and glutamate:

$\text{L-Alanine} + \alpha\text{-Ketoglutarate} \xrightarrow{\text{GPT}} \text{Pyruvate} + \text{L-Glutamate}$

The pyruvate formed is reduced to lactate by lactate dehydrogenase (LDH) with simultaneous oxidation of NADH to NAD⁺:

$\text{Pyruvate} + \text{NADH} + \text{H}^+ \xrightarrow{\text{LDH}} \text{Lactate} + \text{NAD}^+$

The rate of decrease in absorbance at 340 nm due to consumption of NADH is directly proportional to GPT activity in the sample. This is the UV-kinetic (IFCC) method.

Reagents

• Reagent 1 (Buffer/Enzyme): Tris buffer (pH 7.5), L-alanine, LDH, NADH, stabilizers.
• Reagent 2 (Substrate): α-Ketoglutarate.
• Sample: Non-haemolysed serum or plasma (heparin). Avoid EDTA, citrate, oxalate.

Principle of Measurement

Absorbance is measured at 340 nm at 37 °C. The decrease in absorbance per minute (ΔA/min) is proportional to GPT activity.

Procedure

1. Bring reagents and samples to room temperature; reconstitute reagents as per the kit insert.
2. Set the spectrophotometer/analyser at 340 nm, temperature 37 °C.
3. Pipette into a clean dry cuvette:

4. Mix gently and incubate at 37 °C for 1–2 minutes (lag/equilibration phase).
5. Add 0.25 mL Reagent 2 (α-ketoglutarate), mix and immediately start the stopwatch (this initiates the reaction).
6. Read the absorbance at 340 nm every 30 seconds for 3 minutes (kinetic phase).
7. Record the change in absorbance per minute (ΔA/min) during the linear phase.

Calculation

$\text{GPT activity (U/L)} = \frac{\Delta A/\text{min} \times V_t \times 10^6}{\varepsilon \times V_s \times d}$

where:

• $V_t$ = total reaction volume (mL)
• $V_s$ = sample volume (mL)
• $\varepsilon$ = molar extinction coefficient of NADH at 340 nm = $6.22 \times 10^3\ \text{L·mol}^{-1}\text{·cm}^{-1}$
• $d$ = light path length (cm)
• $10^6$ = conversion factor from mol/L/min to µmol/min/L (U/L); $V_t$ and $V_s$ must be in the same unit

Worked factor for the volumes above ($V_t = 1.35$ mL, $V_s = 0.1$ mL, $d = 1$ cm):

$\text{Factor} = \frac{1.35 \times 10^6}{6220 \times 0.1 \times 1} \approx 2170$

so $\text{GPT (U/L)} = \Delta A/\text{min} \times 2170$. Commercial kits supply a pre-calibrated factor for their own volumes (for example, 1768 for $V_t = 1.1$ mL and $V_s = 0.1$ mL).

Reference Range (37 °C)

Clinical Significance

• Raised GPT: Viral hepatitis, alcoholic hepatitis, drug-induced liver injury, cirrhosis, obstructive jaundice, myocardial infarction.
• GPT is more liver-specific than AST; ALT rises mainly in hepatocellular damage.
• AST/ALT ratio > 2 suggests alcoholic liver disease; ratio < 1 suggests viral hepatitis.

Precautions

1. Use fresh, non-haemolysed serum (haemolysis releases intracellular ALT → falsely high).
2. Avoid EDTA/citrate/oxalate — they chelate metal ions and inhibit LDH.
3. Maintain strict 37 °C temperature and 340 nm wavelength.
4. Read only in the linear phase of the reaction.
5. Run a calibrator and quality control sample with each batch.

1. Definition (2 marks)

Stain: A stain is a dye or chemical reagent (usually a coloured organic compound) that has an affinity for specific tissue or cellular components and is used to impart colour to them so that they become visible and distinguishable under the microscope.

Staining: Staining is the technique of applying a dye or a combination of dyes to a biological specimen (tissue section, smear, or cell) in order to produce contrast between different structures, thereby making them visible for microscopic examination and diagnosis.

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2. H&E Staining (8 marks)

H&E (Haematoxylin and Eosin) staining is the routine, most widely used differential staining method in histopathology. Haematoxylin stains nuclei blue/purple (basophilic structures) and eosin stains cytoplasm and extracellular matrix pink/red (acidophilic structures).

Principle

• Haematoxylin is a basic dye; it stains acidic structures (chromatin/DNA, RNA in nucleoli and ribosomes) blue-purple.
• Eosin is an acidic dye; it stains basic structures (cytoplasmic proteins, collagen, muscle) pink-red.
• The two dyes together give good nuclear-cytoplasmic contrast.

Requirements

• Reagents: Harris or Mayer's haematoxylin, 1% acid alcohol (differentiator), ammonia water or Scott's tap water (blueing), eosin (0.5–1% aqueous or alcoholic), xylene, graded alcohols (100%, 95%, 70%), DPX mountant.
• Equipment: staining racks/coplin jars, slides with sections, oven, light microscope.

Procedure (steps)

1. Deparaffinization: Place slides in xylene I and xylene II, 2–3 minutes each, to remove wax.
2. Hydration: Pass through absolute alcohol I, absolute alcohol II, 95% alcohol, and 70% alcohol, 1–2 minutes each, then rinse in running tap water.
3. Haematoxylin staining: Immerse in Harris haematoxylin for 5–10 minutes (or Mayer's for 3–5 minutes).
4. Washing: Rinse in running tap water to remove excess dye.
5. Differentiation: Dip in 1% acid alcohol for a few seconds until nuclei are sharply defined and background is pale.
6. Washing: Rinse well in running tap water.
7. Blueing: Place in ammonia water or Scott's tap water until nuclei turn blue.
8. Washing: Rinse in running tap water.
9. Counterstaining: Stain in eosin for 1–2 minutes.
10. Dehydration: Pass through 70%, 95%, absolute alcohol I and absolute alcohol II, 1–2 minutes each (eosin is decolourized if left too long in alcohol).
11. Clearing: Place in xylene I and xylene II, 2–3 minutes each, until sections become transparent.
12. Mounting: Mount with DPX or Canada balsam and apply coverslip; allow to dry and examine under the microscope.

Results

Precautions

• Do not allow sections to dry at any stage (causes artefacts).
• Keep differentiation time short and controlled.
• Change xylene and alcohols regularly; filter haematoxylin before use.
• Ensure complete deparaffinization for proper staining.