Whole blood (volume ≈ 5 litres in adults) consists of plasma (55–60%) and formed elements (40–45%). Formed elements include RBCs (erythrocytes), WBCs (leukocytes), and platelets (thrombocytes). The ratio of RBCs to total blood volume is the haematocrit (PCV) — normally 40–54% in males, 36–48% in females.

Plasma is the liquid portion of unclotted blood containing water (90%), proteins (albumin, globulins, fibrinogen), electrolytes, hormones, glucose, lipids, and waste products. Serum is plasma from which fibrinogen and other clotting factors have been removed after coagulation. Serum is used for biochemical tests (LFT, RFT, serology), while plasma is used for coagulation tests.

Key clinical point: Fibrinogen is the difference between plasma and serum. Plasma is obtained by centrifuging anticoagulated blood; serum by centrifuging clotted blood. Serum contains antibodies, antigens, and most proteins but not clotting factors.

Venipuncture is the standard method for collecting blood from antecubital veins (median cubital, cephalic, basilic). Tourniquet is applied 5–10 cm above puncture site and released as soon as blood flows. Blood is collected in vacuum tubes (Vacutainer) in a specific order (blood culture → citrate → serum → heparin → EDTA → fluoride). Incorrect order of draw can cause cross-contamination of additives.

Finger prick (capillary puncture) uses a lancet on the 3rd or 4th fingertip, after warming to increase blood flow. First drop is wiped away. Used for CBG, HbA1c, malaria smear, and paediatric CBCs. Ear lobe puncture is an older method, avoided in modern practice. Heel prick is used in neonates. Capillary blood differs slightly from venous blood in composition.

Key errors: Milking/squeezing the finger dilutes the sample. Using excessively tight tourniquet >2 min causes haemoconcentration. EDTA tube should be properly filled to exact ratio (8:1 blood:anticoagulant ratio or as per manufacturer’s specification) to prevent cell shrinkage/swelling.

Anticoagulants prevent blood clotting by different mechanisms: EDTA (lavender/purple top) chelates calcium ions, used for CBC, blood grouping, reticulocyte count. Sodium citrate (blue top) also chelates calcium, used for ESR (Westergren: 3.8%, Wintrobe: potassium oxalate+ammonium oxalate) and coagulation studies (3.2% for PT/APTT at 9:1 ratio). Heparin (green top) activates antithrombin III, used for electrolytes, blood gases, and osmolality. Potassium oxalate precipitates calcium; combined with NaF for glucose tubes (grey top).

Preparation of EDTA vials: Disodium EDTA or dipotassium EDTA at 1–2 mg/mL blood. K₂EDTA is preferred (faster dissolving). Liquid EDTA is more reliable than dry EDTA. For 5 mL blood: add 10 mg EDTA. Tubes must be filled correctly (under/over-filling causes errors in CBC).

Double oxalate (Paul-Heller mixture): Ammonium oxalate (1.2 g) + potassium oxalate (0.8 g) per 100 mL. Used for ESR in Wintrobe method. Ammonium oxalate causes RBC swelling; potassium oxalate causes shrinkage — together they balance each other for isotonic effect.

The Neubauer counting chamber (haemocytometer) is a thick glass slide with two ruled areas. Each ruled area is 9 mm² (3×3 mm) with a central 1 mm² large square divided into 25 medium squares, each subdivided into 16 small squares. Depth = 0.1 mm. WBCs counted in 4 corner large squares (4 mm²); RBCs in 5 small squares (4 corners + center of central square). The coverslip is placed so Newton’s rings appear at the edges, confirming correct depth.

Diluting pipettes: Red cell pipette (RBC pipette) — graduated to 0.5 and 1.0, bulb marked 101 (total volume). Dilution: blood to 0.5, diluting fluid to 101 = 1:200 dilution. White cell pipette — graduated to 0.5 and 1.0, bulb marked 11. Dilution: blood to 0.5, fluid to 11 = 1:20 dilution. First 3–4 drops are discarded before loading chamber (dead space contains only diluent).

Sahli’s haemoglobinometer contains a graduated tube, comparator with standard brown glass, N/10 HCl, and a stirring rod. Blood (0.02 mL) is drawn to mark, placed in tube containing 5 drops of N/10 HCl, mixed, and diluted with distilled water until colour matches standard. Reading in g/dL. Bulk dilution uses tubes instead of pipettes — simpler, less precise, used when multiple samples are processed.

A thin blood smear (peripheral blood film) is prepared by placing a small drop of blood near one end of a slide, spreading with a spreader slide at 30–45°, and air-drying rapidly. A good smear has a feather edge (tail), covers 2/3 of slide, no holes, uniform thickness. Used for differential WBC count, RBC morphology, and platelet estimation. Disadvantage: fewer parasites visible per field.

A thick blood smear is prepared by spreading 2–3 drops of blood in a circle (1.5–2 cm diameter) without fixation. Haemoglobin is laked (RBCs lyse) by placing in water before staining. This concentrates WBCs and parasites. Advantage: 20× more sensitive than thin smear for detecting malaria and microfilaria. Disadvantage: RBC morphology cannot be assessed, species identification may be harder.

Key points: Thick smear is not fixed with methanol (fixation prevents laking). Thin smear IS fixed before staining. Both are stained with Giemsa or Leishman. For malaria, the WHO recommends thick smear as the gold standard. Smears should be made within 1 hour of collection if EDTA blood is used (longer time distorts cell morphology).

WBC count — Diluted 1:20 with Turk’s fluid. Counted in 4 large corner squares of Neubauer chamber. Formula: WBC/µL = N × 20 × 10 / 4 = N × 50 (where N = cells in 4 squares). Normal: 4,000–11,000/µL. Leukocytosis (>11,000): bacterial infection, inflammation, leukaemia, steroids. Leukopenia (<4,000): viral infections, aplastic anaemia, SLE, cytotoxic drugs.

RBC count — Diluted 1:200 with Hayem’s or formal-citrate solution. Counted in 5 small squares (center area of Neubauer chamber). Formula: RBC/µL = N × 200 × 10 / (5 × 1/25) = N × 10,000. Normal: males 4.5–5.5 million/µL; females 3.8–5.1 million/µL. Low: anaemia; High: polycythaemia, dehydration, high altitude.

Platelet count (indirect method — Fonio’s method): count on stained smear; direct method uses Rees-Ecker fluid (1:100). Normal: 1.5–4.5 × 10⁵/µL. Thrombocytopenia (<1,50,000): dengue, ITP, aplastic anaemia, chemotherapy — bleeding risk increases when <50,000. Thrombocytosis (>4,50,000): reactive (infection, post-splenectomy) or essential thrombocythaemia.

Pre-analytical errors are the most common (60–70% of all lab errors): wrong anticoagulant or ratio, improper mixing, clotted sample, haemolysis, lipemic sample, delay in processing, incorrect patient ID, prolonged tourniquet application, squeezing the finger-prick site.

Analytical errors: improper dilution (pipetting error), uneven distribution in chamber, incorrect volume loaded, dirty or cracked chamber, air bubbles under coverslip, not discarding first drops from pipette, counting cells touching top/bottom borders differently, instrumental drift, poor calibration.

Post-analytical errors: mathematical calculation errors, transcription errors, incorrect reference range applied, reporting to wrong patient. Quality control measures include daily controls, Levey-Jennings charts, Westgard rules, and regular instrument calibration. Duplicate counts help estimate random errors (CV should be <5%).

Differential count identifies 5 main WBC types in 100 consecutive leukocytes on a stained smear. Neutrophils (50–70%): multilobed nucleus (2–5 lobes), pink granules, 10–14 µm. Lymphocytes (20–40%): large round dark nucleus, scant blue cytoplasm, 7–12 µm (small). Monocytes (2–8%): largest WBC, kidney/horseshoe nucleus, grey-blue cytoplasm, fine azurophilic granules, vacuoles. Eosinophils (1–4%): bilobed nucleus, large coarse orange granules. Basophils (0–1%): S-shaped nucleus obscured by coarse blue-black granules.

Clinical significance: Neutrophilia — bacterial infection; Lymphocytosis — viral infections (EBV, CMV), CLL; Monocytosis — TB, SBE, recovery phase; Eosinophilia — allergy, parasites; Basophilia — CML, polycythaemia. Band cells (immature neutrophils) >6% = left shift indicating acute infection/sepsis.

The battlement technique is used to systematically scan the smear (edge to edge, in a zigzag pattern at the junction of the thin and thick area) to avoid differential counting bias. 100 cells minimum; 200 cells for greater accuracy.

ESR (Erythrocyte Sedimentation Rate) measures how fast RBCs settle in a standing tube over 1 hour. It depends on rouleaux formation (RBCs stacking like coins) promoted by fibrinogen, globulins, and acute-phase proteins. Three phases: Rouleaux formation (10 min), fast sedimentation (40 min), packing phase (10 min).

Westergren method (ICSH recommended): Blood:3.8% sodium citrate = 4:1. Tube: 300 mm long, 2.5 mm bore. Set vertically at room temperature. Read at 1 hour. Normal: males <15 mm/hr; females <20 mm/hr. Wintrobe method: Undiluted blood (with oxalate). Tube: 110 mm, 2.5–3 mm bore. Same reading time. Normal: males 0–9 mm/hr; females 0–20 mm/hr.

Elevated ESR: TB, RA, SLE, infections, malignancy, pregnancy, anaemia, multiple myeloma. Low ESR: polycythaemia, sickle cell, spherocytosis, hypofibrinogenaemia. ESR is a non-specific test (acute phase reactant). Westergren is more sensitive and reproducible; preferred internationally.

Haemoglobin estimation methods include: (1) Cyanmethaemoglobin method (HiCN) — WHO reference method. Blood is added to Drabkin’s solution, converting all Hb (except SHb) to stable cyanmethaemoglobin. Absorbance read at 540 nm; compared with standard curve. Accurate and reproducible. (2) Sahli’s method — N/10 HCl converts Hb to acid haematin; compared with amber glass comparator. Simple, cheap, but inaccurate. (3) Oxyhaemoglobin method — diluted in N/10 NH₄OH, read at 540 nm. (4) Haemoglobin-meter (portable) — POCT devices.

Preparation of Standard Curve (HiCN method): Use certified cyanmethaemoglobin standard (e.g., 80 mg/dL cyanmetHb = 20 g/dL Hb). Prepare serial dilutions (e.g., 100%, 75%, 50%, 25%, 0%). Measure absorbance of each at 540 nm. Plot absorbance (Y-axis) vs Hb concentration (X-axis). Draw best-fit line. Read unknown samples from the curve.

Normal Hb: males 13.5–17.5 g/dL, females 12–16 g/dL, children 11–13 g/dL. Anaemia: WHO defines as Hb <13 g/dL (males), <12 g/dL (females). Severe anaemia <8 g/dL. Causes: IDA, B12/folate deficiency, haemolysis, chronic disease, blood loss.

Drabkin’s solution is the reagent used for cyanmethaemoglobin (HiCN) method. Composition (per 1 litre): Potassium ferricyanide [K₃Fe(CN)₆] — 200 mg; Potassium cyanide [KCN] — 50 mg; Sodium bicarbonate [NaHCO₃] — 1,000 mg; Distilled water — up to 1 litre. Some formulations add 0.5 mL Sterox SE (detergent) to prevent turbidity from lipids.

Principle: K₃Fe(CN)₆ oxidises haemoglobin to methaemoglobin (Fe²⁺→Fe³⁺). KCN then converts methaemoglobin to stable cyanmethaemoglobin (HiCN), which has a stable absorption peak at 540 nm. NaHCO₃ maintains pH. Ratio: 0.02 mL blood in 5 mL Drabkin’s = 1:250 dilution. Mix and wait 3–5 min. Read at 540 nm.

Safety precautions: KCN is toxic — avoid contact with acids (releases HCN gas). Dispose waste in alkaline solution. Store in dark bottle (light-sensitive). Shelf life: 6 months if stored at 4°C. Solution must be pale yellow; discard if colour changes. Standard certified Hb solution must accompany each batch.

Romanowsky stains are polychromatic stains composed of methylene blue derivatives (azure B) + eosin, dissolved in methanol (acts as both fixative and solvent). They produce a characteristic purple colour called “Romanowsky effect” — an artefact arising from interaction of methylene azure and eosin. Leishman stain: ready-to-use, air-dried smear stained for 15 sec (fixes), then diluted with buffer for 15 min. Easiest to use. Wright stain: similar to Leishman, popular in USA. Giemsa stain: best for malaria/parasites; diluted 1:10 with phosphate buffer (pH 7.2), applied for 20–30 min on water-laked thick smears.

Staining results: Nuclei → purple/violet; cytoplasm of RBCs → pink; neutrophil granules → lilac/pink; eosinophil granules → orange-red; basophil granules → deep blue/purple; lymphocyte cytoplasm → sky blue; monocyte cytoplasm → grey-blue. Parasites (malaria): chromatin (nucleus) → red; cytoplasm → blue.

Preparation of Leishman stock solution: Dissolve 0.15 g Leishman powder in 100 mL acetone-free methanol (with glass beads, shake daily for 3 days). Filter and store in dark amber bottle. Stable for months. Working solution: dilute 1:1 with phosphate buffer (pH 6.8–7.0) at time of use. Correct pH is critical: acidic pH (↓) → RBCs too red, WBC nuclei not visible; alkaline pH (↑) → everything blue.

N/10 HCl (0.1 N HCl = 0.1 M HCl) is used in Sahli’s method to convert haemoglobin to brown acid haematin. Preparation: Concentrated HCl is 36–38% pure with specific gravity 1.19. Normality of concentrated HCl ≈ 11.3 N. To prepare 1 litre of 0.1 N HCl: Volume required = (0.1 × 1000) / 11.3 = 8.85 mL. Add 8.85 mL conc. HCl to about 900 mL distilled water, then make up to 1 litre (always add acid to water — never reverse).

Alternatively, use the formula: N₁V₁ = N₂V₂ where N₁ = normality of concentrated HCl, V₁ = volume to be taken, N₂ = 0.1 N, V₂ = 1000 mL. Standardise with sodium carbonate primary standard if precision is required. Store in glass-stoppered bottle; HCl is volatile and corrosive.

Safety: Always add acid to water (exothermic reaction). Use fume hood. Wear gloves and goggles. Concentrated HCl fumes are corrosive to respiratory tract. Label clearly: “0.1 N HCl — CORROSIVE.” Check concentration periodically; HCl concentration decreases on storage.

Bleeding Time (BT) tests primary haemostasis (platelet plug formation + vascular response). Duke’s method (earlobe): lancet puncture 3 mm deep; blot with filter paper every 30 sec; BT = time from puncture to cessation of bleeding. Normal: 1–3 min. Ivy’s method (forearm): BP cuff at 40 mmHg, two incisions 1 mm deep × 5 mm long; Normal: 2–7 min. Prolonged BT: thrombocytopenia (<100,000), platelet dysfunction (VWD, aspirin, uraemia), vascular disorders (scurvy).

Clotting Time (CT) tests secondary haemostasis (intrinsic coagulation pathway). Lee-White method: venous blood in 3 glass tubes at 37°C. CT = time for clot formation in last tube. Normal: 5–11 min (whole blood). Prolonged CT: haemophilia A and B, severe factor deficiencies, heparin therapy, DIC, severe liver disease.

Important: BT and CT are screening tests only and have largely been replaced by PFA-100 (for BT) and PT/APTT (for CT) in modern labs. BT is not affected by coagulation factor deficiencies (only by platelets and vessels). CT does not detect mild factor deficiencies. Both tests have high intra/inter-observer variability.

Malaria diagnosis: Giemsa-stained thick and thin smears are gold standard. Species differentiation: P. vivax — enlarged RBC, Schüffner’s dots, amoeboid trophozoite, 16-cell schizonts. P. falciparum — small ring forms (double dots/appliqué), multiply infected cells, banana-shaped gametocytes, no enlarged RBC, Maurer’s clefts. P. malariae — “band form” trophozoite, 8-cell rosette schizont, Ziemann’s dots. P. ovale — oval/fimbriated RBC, Schüffner’s dots, 8-cell schizonts.

Microfilaria is the larval stage of filarial worms (Wuchereria bancrofti, Brugia malayi, Loa loa). In blood smear: sheathed (W. bancrofti, B. malayi) or unsheathed (Mansonella); identified by sheath, nuclear column, and tail morphology. W. bancrofti: sheathed, no nuclei in tail tip. B. malayi: sheathed, 2 distinct nuclei in tail tip. Blood collected at midnight (nocturnal periodicity for W. bancrofti).

Thick smear technique for microfilaria: spread thick drop, allow to dry without fixing, dehemoglobinise with water, stain with Giemsa. Membrane filtration (Nucleopore filter) concentrates microfilaria from 1–5 mL blood — more sensitive. Modified Knott’s technique: 1 mL blood + 9 mL 2% formalin, centrifuge, stain sediment — detects even low microfilaraemia.

The ABO blood group system is based on the presence of A and B antigens on RBC surface and corresponding antibodies (isoagglutinins) in plasma. Group A: A antigen, anti-B antibody. Group B: B antigen, anti-A. Group AB: both antigens, no antibodies (universal recipient). Group O: no antigens, anti-A and anti-B (universal donor). Forward grouping tests RBCs with known antisera (anti-A, anti-B); Reverse grouping tests serum with known A and B cells. Both must match.

Rh typing: D antigen on RBC = Rh positive (85% of population). Test: mix RBCs with commercial anti-D serum (IgM) on slide/tube. Agglutination = Rh positive; no agglutination = Rh negative. Clinical significance: Rh incompatibility in pregnancy causes haemolytic disease of the newborn (HDN) — Rh-ve mother, Rh+ve fetus; anti-D antibodies formed, cross placenta in subsequent pregnancies. Prevention: anti-D immunoglobulin (Rho-GAM) within 72 hours post-delivery.

Sources of error in blood grouping: rouleaux mimicking agglutination, cold autoagglutinins, polyagglutinability, incorrect reagents, mislabelled samples, incorrect centrifugation. The slide method is quick but less accurate; tube method is more reliable. Crossmatching (compatibility test) is done before transfusion.

Quality control (QC) in haematology ensures accuracy and precision of test results. Internal QC (IQC): daily use of commercial control materials (assayed and unassayed) at 2–3 concentration levels. Results plotted on Levey-Jennings chart against mean ± 2SD and ± 3SD limits. Westgard rules detect systematic (1₂s warning, 2₂s, 4₁s) and random (1₃s, R₄s) errors. External QA (EQA): participation in national/international proficiency testing programmes (e.g., WHO EQAS, NEQAS).

Key QC parameters: Accuracy = closeness to true value; Precision = reproducibility of repeated measurements; CV% = (SD/mean) × 100 — acceptable CV for Hb: <2%; WBC: <5%; platelets: <5%. Bias = systematic difference from true value. Important: control materials should bracket the patient values (low, normal, high ranges).

Calibration of haematology analysers uses manufacturer-provided calibrators or fresh normal blood with assigned values. Daily: run QC before patient samples; check reagent and instrument status; check reagent expiry. Weekly: check diluter function, temperature, carry-over. Monthly: verify reference ranges, review QC statistics. Delta check: compare current with previous result for same patient to detect errors.