Nematology

Introduction

Nematology is a branch of biological science dealing with nematodes, a diverse group of microscopic, triploblastic, bilaterally symmetrical, unsegmented roundworms with a pseudocoelom.

Habitat: Found worldwide — from Arctic to tropics, ocean depths to mountaintops, and especially abundant in soil.

Abundance: Comprise 80–90% of all multicellular animals.

Types & Roles : Known as eelworms, roundworms, nemas.

Categories:

•  Plant parasitic nematodes (phytonematodes) – attack crops.

•  Animal/human parasites (helminths) – studied under Helminthology.

•  Free-living and beneficial species – contribute to soil health.

•  The word “nematode” originates from Greek: nema (thread) + eidos (form).

Impact on Crops

• Attack roots, stems, buds, leaves, seeds.

• Cause symptoms like stunting, yellowing, slow growth, often mistaken for nutrient deficiency.

Nematode Distribution

•   Sea water: 50%

•   Free-living: 25%

•   Animal parasites: 15%

•   Plant parasites: 10%

History of Nematology

I. Early History (1743–1940)

1743 – Needham, T.: First record of plant parasitic nematode (Anguina tritici) in wheat.

1855 – Berkeley, M.J.: Discovered root-knot nematode (Meloidogyne spp.).

1857 – Kuhn, J.: Described stem nematode (Ditylenchus dipsaci).

1859 – Schacht, H.: First reported cyst-forming nematode in sugar beet.

1871 – Schmidt, A.: Described Heterodera schachtii.

1891 – Ritzema Bos, J.: Discovered foliar nematodes (Aphelenchoides spp.).

1914–1932 – Cobb, N.A.: Father of American Nematology; developed techniques for sampling, mounting, and identifying nematodes.

1930 – Filipjev, I.N.: Published agricultural nematode book.

1937 – Chitwood, B.G.: Wrote Introduction to Nematology.

II. New Era in Nematology (1941–1990)

1941 – Cannon, O.S.: Reported Heterodera rostochiensis (Golden Nematode).

1944 – Christie & Albin: Identified races of root-knot nematodes.

1949 – Chitwood, B.G.: Described Meloidogyne species; developed host range and ID keys.

1953 – Christie & Perry: Highlighted role of ectoparasitic nematodes.

1955 – Mountain, W.B.: Cultured nematodes in sterile conditions.

1958 – Hewitt, W.B.: Found nematode role in virus transmission.

III. Nematology in India

1901 – Barber, C.A.: First report of root-knot in tea (South India).

1934 – Ayyar, P.N.K.: Root-knot in vegetables.

1936 – Dastur, J.F.: White tip in rice (Aphelenchoides spp.).

1961 – Jones, F.G.W.: Potato cyst (Globodera rostochiensis) in Nilgiris.

1965: Radopholus similis in banana (Kerala).

1966: Division of Nematology established at IARI, New Delhi.

1971: Launch of Indian Journal of Nematology.

1977: AICRP on Nematode Pests (14 centers).

Scope of Nematology in Agriculture :

•   Crop Protection: Manages plant parasitic nematodes (Meloidogyne, Heterodera, Radopholus), reducing crop losses.

•   Resistant Varieties: Aids in developing nematode-resistant crops through breeding and biotechnology.

•   Integrated Management: Supports eco-friendly control via cultural, biological, chemical, and physical methods.

•   Soil Health: Uses free-living nematodes as indicators of soil quality and fertility.

•   Diagnostics & Forecasting: Enables early detection, pest forecasting, and site-specific nematode control.

Nematode – Morphology

Nematodes are triploblastic, bilaterally symmetrical, unsegmented, colourless,    pseudocoelomate, and vermiform animals with a circular cross-section. 

When relaxed with gentle heat, they may lie

●       Straight (e.g., Pratylenchus)

●       Sightly curved (Hoplolaimus),

●       C-shaped (Tylenchorhynchus),

●       Spiral (Helicotylenchus). 

Some exhibit sexual dimorphism, and females may take on lemon, pear, kidney, or saccate shapes.  

Size & Body Regions:

 Nematodes range from 0.2 mm to 12 mm in length, averaging 0.01–0.5 mm in breadth. Males are typically smaller than females

 The body is divided into regions:

Ø  Head (with mouth, lips, stoma),

Ø  Neck (to oesophagus),

Ø  Tail (post-anal region).

1. Anterior Region (Head):

Contains mouth, stylet (in plant parasites), lips, amphids, and pharynx. Functions in feeding, host penetration, and sensory perception.

2. Middle Region (Body/Trunk):

Houses oesophagus, intestine, reproductive organs, nerve ring, and excretory system. Involved in digestion, reproduction, and coordination.

3. Posterior Region (Tail):

Includes anus, cloaca (in males), spicules (in males), and phasmids (in some). Functions in excretion, mating, and sensory detection.

The body is further divided longitudinally into ventral, dorsal, right, and left laterals.

General Structure:

Nematode bodies are tubular and divided into:

1. Outer body tube – Body wall

2. Inner body tube – Digestive system

3. Body cavity – Reproductive, nervous, and excretory systems

 Outer Body Tube Includes: Exoskeleton, Hypodermis, Muscle layer

A. Cuticle (Exoskeleton): Tough, non-cellular, semipermeable layer secreted by hypodermis; covers body and all natural openings. Often features taxonomically important surface markings.

1. Cuticular Lining:

a. Punctations: Small, round structures aiding cuticle strength and protein transport.

b. Annules/Striations: Transverse lines giving a segmented appearance (e.g., in Criconemoides, root-knot nematodes); essential for movement and species identification.

c. Cuticular Markings (Longitudinal):

1. Ridges: Raised lines running along the body (sub-median/lateral surfaces).

2. Alae: Lateral or sub-lateral projections aiding in locomotion:

3. Caudal alae: Found in males’ posterior (copulatory bursa).

4. Cervical alae: Present in the anterior body (some marine species).

5. Longitudinal alae: Confined to lateral fields; help in movement and minor width change

2.Cuticle Structure (Three Layers):

a) Cortical layer: Thin outermost layer (25–40 mµ), proteinaceous (keratin-

like); forms a tough cyst in females of cyst nematodes.

b) Median layer: ~0.1 µ thick; protein similar to collagen, offers flexibility.

c)  Basal layer: Thickest (125–500 mµ), with strong protein bonds; provides environmental resistance

Cuticle Functions:

•   Protects from harsh conditions

•   Acts as an exoskeleton

•   Aids in movement through soil and plant tissue

B. Hypodermis:

A metabolically active layer under the cuticle, forms four cords (dorsal, ventral, two lateral) and contains glands.

C. Muscle Layer:

Single layer of spindle-shaped cells attached to the hypodermis. Controlled by dorsal/ventral nerves, causing sinusoidal movement.

Nemotode Anatamy

Nematode Digestive System

The gut is a straight tube, divided into three main regions:

1. Stomodaeum (Foregut)

Mouth & Lips: 6 lips (may fuse); used in feeding.

Stoma (Buccal Cavity): May have teeth or stylet:

Stomatostylet (with knob, e.g. Tylenchida)

Odontostylet (no knob, e.g. Dorylaimida)

Oesophagus (Pharynx): Muscular pump with:

Corpus – contains glands, nerves, muscles

Isthmus – narrow connecting tube

Basal Bulb – has valves to prevent backflow

2. Mesenteron (Midgut)

Straight endodermal tube made of epithelial cells.

Regions: Anterior (ventricular), Mid-intestine, Posterior (pre-rectal)

3. Proctodeum (Hindgut)

Females: Rectum + slit-like anus, controlled by H-shaped depressor muscle.

Males: Cloaca – common outlet for digestive and reproductive tracts; contains spicules and copulatory structures

Pharyngeal (Esophageal) Glands

Three uninucleate glands: 1 dorsal, 2 subventral

Connected to: Oesophagus via terminal ampulla

Functions: Aid in egg hatching,Host tissue penetration,Digestion

Rectal Glands

Vary by species and sex

Secrete: Gelatinous mucopolysaccharide matrix around egg masses.

Function: Protects egg

Body cavity / Pseudocoelom

 Not a true coelom (not completely mesoderm-lined)

a) Coelomic Cavity (True coelom):

Outer lining: Somatic muscles (mesodermal)

Inner lining: Alimentary canal (ectodermal)

b) Pseudocoelomic Cavity:

Lined with mesodermal tissues

Contains pseudocoelomic fluid

c) Pseudocoelomic Fluid:

Bathes internal organs

Composition: Proteins, glucose, Na⁺, K⁺, Mg²⁺, Zn²⁺, Fe, Cl⁻, P, Cu, hematin, ascorbic acid

Functions: Supports reproductive, nervous, and excretory systems

Note: Circulatory & respiratory systems absent

Reproductive System in Nematodes

General:

Dioecious: Males and females separate; males are smaller and fewer.

Some species show hermaphroditism or parthenogenesis (absence of males).

Female Reproductive System:

Ovary Types:

•   Monodelphic: One ovary

•   Didelphic: Two ovaries

•   Prodelphic: Ovary directed anterior to vulva

•   Opisthodelphic: Ovary directed posterior to vulva

•   Amphidelphic: Two ovaries, one anterior & one posterior to vulva

1. Ovary:

•   Hollow, elongate tube with a cap cell (germinal zone)

•   Followed by growth zone with enlarging oocytes in rows

2. Oviduct:Transports oocytes; may function as spermatheca

3. Uterus:Largest gonad section,Site of fertilization, eggshell formation, egg laying,May also act as spermatheca

4. Vagina:Short, narrow, cuticle-lined tube; connects uterus to vulva

5. Vulva:External female gonopore; usually mid-body

Male Reproductive System:

1. Testes Types: Monarchic: One testis Diarchic: Two testes

2. Testis: Germinal zone (spermatogonia) → growth zone (spermatocytes)

3. Vas deferens: Anterior glandular, posterior muscular region

4. Ejaculatory duct: Opens into cloaca; expels sperm. Associated with copulatory structures (spicules, gubernaculum)

Excretory System in Nematodes

Not well developed, Excretory pore: Mid-ventral, near nerve ring

Types:

Glandular Type (Adenophorea):

Single renette cell

Renette → ventral gland → duct + ampulla → pore

Tubular Type (Secernentea):

Four cuticularised canals (2 anterior, 2 posterior)

Lateral canals join at central pouch → excretory pore

Tubular Subtypes:

•   Asymmetrical/Tylenchid type

•   Inverted 'U'/Ascarid type,,

•   Rhabditid type

•   Simple ‘H’/Oxyurid type

1. Central Nervous System (CNS)

Nerve ring (circum-oesophageal commissure) surrounds oesophagus.

Tylenchida: encircles isthmus, Dorylaimida: encircles anterior oesophagus

Contains 6 papillary ganglia: 2 dorsal, 2 ventral, 2 lateral

Transverse commissures connect longitudinal nerves across body regions

2. Peripheral Nervous System (PNS)

A. Somatic Nerves (run through hypodermis)

Dorsal somatic nerve – from dorsal ganglia to anal region

Latero-dorsal & latero-ventral nerves – paired, submedian

Ventral nerve – part of CNS

Lateral & dorso-lateral nerves – near anal region, with lumbar ganglia

B. Cephalic Papillae Nerves

Originate from cephalic ganglia, extend near lips

C. Amphidial Nerves

Linked via sub-ventral trunk, end in amphid glands

Path: Aperture → Fovea → Canalis amphidialis → Sensilla pouch → Amphidial nerve

3. Sensory Structures

Amphids: Paired chemoreceptors in head; variable aperture shapes; may differ by sex

Deirids: Paired mechanoreceptors near excretory pore (no external opening)

Phasmids: Posterior sensory organs near tail; may be enlarged as scutella

Caudalids: Nerve commissure linking pre-anal to lumbar ganglia

Cephalids: Refractive rings behind head; origin of lateral cords

Techniques of Extraction Of Nematode

Collection of soil and root samples for nematode extraction:

Sampling from field crops:

❖      Leave 1 m peripheral area of the field.

❖      Remove top 2–3 cm soil using hand hoe.

❖      Collect 200 cc soil with feeder roots from 15–20 cm depth (subsample).

❖      Take 10–20 subsamples per hectare in a zigzag pattern.

❖      Combine subsamples into one polythene bag (composite sample).

❖      Reduce composite sample by quartering for a representative sample.

❖      Label with sample number/details using aluminium foil or paper, place in bag, and tie with rubber band.

Vegetable Crop

❖      Select 6 rows (2 each from start, middle, end).

❖      Collect 8–10 subsamples (20–30 cm depth) per row pair in zigzag.

❖      Combine in one polythene bag and label.

❖      Reduce by quartering (representative sample).

Orchard

❖      Take 2 subsamples/tree (30–60 cm depth, feeder root zone).

❖      Sample 10 trees randomly per hectare.

❖      Pool in one polythene bag, label, and quarter.

Tree

❖      Collect 5 subsamples each from main stem and drip line.Depth

❖      varies with tree type and age.Combine in one polythene bag, label.

1.Extraction of vermiform nematodes:

Cobb’s Decanting and Sieving Method:

Principle:

Nematodes and soil particles settle at different rates due to specific gravity differences.

Procedure:

❖      Mix composite soil sample; take 250 cc in Pan I.

❖      Add 1 L water, stir to break clods.

❖      After 30 sec, decant through 20-mesh sieve into Pan II (repeat twice). Discard residue in Pan I.

❖      Stir Pan II gently, wait 30 sec, then sieve through 60, 100, 200, and 350 mesh sieves.

❖      Discard filtrate from 350 mesh.

❖      Collect and label residues from 60, 100, and 200 mesh in separate beakers.

❖      Use 60-mesh residue for cyst nematode examination.

❖      Further process 100, 200, and 350-mesh residues for other nematodes.

 Baermann’s Funnel Method :

 Principle:

 Active nematodes migrate through tissue paper into water due to gravity, 

 while soil debris stays on top.

 Procedure:

❖      Pre-process soil using 20, 60, 100, 200, 350 mesh sieves.

❖      Set up a 10 cm glass funnel with rubber tube closed by a Hoffman’s clip.

❖      Fill the funnel with water. Place double-layered tissue paper on a wire gauze and set it in the funnel.

❖      Let Cobb’s suspension settle; decant and pour the remaining onto the tissue paper. Ensure gauze just touches the water surface.

❖      Allow nematodes to migrate into water and collect at the tube base.

❖      After 24–48 hours, release the clip and collect nematode suspension.

Modified Baermann’s Funnel Technique:

Procedure:

❖      Place double-layered tissue paper on a mesh or support inside a Petri dish.Add nematode suspension (from Cobb’s method) on the tissue paper.

❖      Pour water so the bottom of the tissue touches the water surface.Active nematodes migrate through the tissue into water.

❖      Collect nematodes from the Petri dish after 24–48 hours.

Centrifugal Flotation Method

Principle:

Nematodes (specific gravity 1.05) float in sugar solution (1.18) during centrifugation, enabling separation.

Sugar Solution Prep:

Dissolve 484 g sugar in water to make 1 L. Add 10% lactic acid to prevent microbial growth.

Procedure:

❖      Prepare soil suspension using Cobb’s method.

❖      Pour suspension into centrifuge tubes (equal volume, leave 0.5 cm space).

❖      Centrifuge at 3500 rpm for 3 min; discard supernatant.

❖      Add sugar solution (half tube), shake, top up to 0.5 cm from top.

❖      Centrifuge again at 3500 rpm for 1 min. Decant supernatant through 350-mesh sieve.

❖      Wash residue on sieve with water to remove sugar. Transfer cleaned residue to a beaker for nematode observation.

2.Extraction of cysts from soil samples:

Conical Flask Method

Principle:

Dry cysts, being lighter, float in water and adhere to the container walls due to surface tension.

Procedure:

❖      Shade-dry the soil sample.

❖      Add a known amount of soil to a conical flask and fill with water.

❖      Shake thoroughly and top up to the rim.Let it stand for 10 minutes.

❖      Floating cysts adhere near the rim.

❖      Pour the suspension through filter paper placed over a beaker.Cysts remain on the filter paper.

❖      Dry and examine the filter paper under a microscope.

Fenwick Can Method

Principle:

Dry cysts, being lighter, float in water and adhere to the container walls due to surface tension.

Procedure:

❖      Fill the can with water and place shade-dried soil in the 20 mesh sieve.Wash the soil sample with water.

❖      Cysts pass into the collar and are trapped in the 60 mesh sieve. Allow water to drain.

❖      Wash sieve contents into a white enamel basin.

❖      Collect floating cysts from the basin’s periphery

3.Extraction of nematodes from roots :

Direct Examination

Procedure:

❖      Wash and chop infested plant material.Place in Petri dish with water.

❖      Migratory and endo/semi-endoparasitic nematodes migrate into water and are visible under a microscope.

❖      Alternatively, process with modified Baermann’s funnel.

Root Incubation Method

Principle:

Nematodes exit roots due to suffocation.

Procedure:

❖      Wash roots and make longitudinal cuts.Place in a jar/polythene bag and loosely seal.

❖      Incubate at 15°C for 72 hours.

❖      Flush roots thrice with water.

❖      Pass wash through 350-mesh sieve and collect residue in a beaker.

Mechanical Maceration / Homogenizer Technique

Principle:

Maceration dislodges nematodes from root tissues by mechanical force.

Procedure:

❖      Wash and chop roots (0.5–1.0 cm).   Blend with water for 15 sec (30–60 sec for hard roots).

❖      Filter through 60-mesh then 350-mesh sieve.

❖      Discard coarse residue; collect nematode suspension on 350-mesh sieve.

❖      Transfer suspension to modified Baermann’s funnel.

Staining Nematodes in Roots

Procedure:

❖      Wash roots, blot dry, cut into 1 cm bits.Boil in acid fuchsin lactophenol for staining.

❖      Drain stain, rinse roots with water.Submerge in plain lactophenol overnight for destaining.

❖      Observe under microscope; nematodes appear pink/red inside transparent roots

Preservation of Nematodes

Killing of Nematodes:

Small Samples:

Transfer a few nematodes to a drop of water on a slide.Kill them by  Holding the slide over a flame for 5–6 seconds, or Adding 2 drops of hot water.

Large Samples:

Concentrate suspension in a vial. Immerse vial in hot water (60°C) for 2 minutes.

Fixing of Nematodes

⮚      Mix equal quantity of hot fixative with concentrated nematode suspension in a screw-cap vial.Label vial with: Nematode name, Collection source, Fixative used, Collection and fixation dates

⮚      The vial with fixed nematodes is called a wet collection.

Processing

⮚      Processing of nematode specimens clears the internal body contents The fixed nematodes and makes clearly visible. Specimens can be processed

⮚      With lactophenol, lactoglycerol or glycerol, which also serve as suitable

⮚      Mountant.

Lactophenol Method (Quick clearing & mounting)

Composition: Phenol: 1 | Lactic acid: 1 | Glycerol: 2 | Water: 1

Heat lactophenol to 65°C on hot plate.Transfer fixed nematodes to hot solution, leave for 2–3 min.

Slow Glycerol Method (Long-term clearing)

Place specimens in 2 ml of 1.5% glycerol in water.Add picric acid or copper sulphate to prevent mold.Keep in desiccator for ~4 weeks for gradual dehydration.

Rapid Seinhorst’s Method (Faster glycerin processing)

Seinhorst Solution I: 96% Ethanol – 20 ml, Glycerol – 1 ml,Distilled water – 79 ml

Seinhorst Solution II: Glycerol – 5 parts, 96% Ethanol – 95 parts

Steps:

⮚      Place fixed specimens in cavity block with 1/3 volume of Solution I.Keep cavity block in a desiccator over 96% ethanol for 12 hrs.Remove excess ethanol without disturbing specimens.

⮚      Add Solution II, leave lid slightly open.Place in desiccator with calcium chloride at room temp.

⮚      After 3 days, transfer specimens in pure glycerin for mounting.

Baker’s Method:

Place Baker’s Solutions I–V in respective cavities on Baker’s slide.Keep slide in oven at 55°C.Sequentially transfer nematode specimens from Solution I to V, allowing 10 minutes in each.

Mounting of Nematodes:

Temporary Mounting

1.Water Mount

⮚      Drop of water on glass slide (76 × 25 mm).

⮚      Transfer nematodes and cover with 1.9 cm cover slip.

⮚      Seal edges with glyceel/nail polish.

2.Fixative Mount

Use a drop of fixative instead of water.

Semipermanent Mounting

⮚      Place nematodes in warm lactophenol, heat for 3–4 sec. Transfer to dehydrated glycerine.

⮚      Arrange 8 nematodes in 2 rows and 3 glass rods in triangle.

⮚      Apply cover slip, drain excess fluid, seal with nail polish, and label

Permanent Mounting (Cobb’s Slide Method)

⮚      Mount in glycerine on a square cover slip (25 × 25 mm).Add nematodes + 3 glass wool pieces in triangle position.

⮚      Cover with 18 mm round cover slip, seal with glyceel/nail polish/paraffin wax.

⮚      Insert into Cobb’s aluminum holder, place two cardboard spacers on sides.

Entomopathogenic Nematodes

EPNs are beneficial nematodes that parasitize crop insects, especially lepidopterans and coleopterans. They act as biopesticide.

Typical Characters of EPNs

⮚      Active host-seeking via chemoreceptors in soil and on plants.

⮚      Quick insect kill by releasing symbiotic bacteria.

⮚      Broad host range: Coleoptera, Lepidoptera, Diptera, Orthoptera, Homoptera.

⮚      Easily mass-multiplied; recoverable from soil.

⮚      Culturable on artificial diet or live hosts; long shelf life in various states.

⮚      Compatible with pesticides; usable as dusts, sprays, capsules, granules, or via irrigation.

⮚      Safe to vertebrates, plants, and non-targets.

⮚      Eco-friendly, self-perpetuating, and registration-exempt.

⮚      Belong to genera: Steinernema, Neosteinernema, Heterorhabditis.