Unit 3: Aseptic culture establishment - Subjective Questions

Aseptic technique refers to the set of practices and procedures performed to prevent contamination by unwanted microorganisms (bacteria, fungi, yeasts) during the establishment and maintenance of plant tissue cultures.

Importance:

• Plant tissue culture media are rich in nutrients (sugars, vitamins, minerals) which support rapid growth of microbial contaminants.
• Microbes grow much faster than plant cells, quickly overtaking and killing the explant.
• Contamination leads to competition for nutrients, release of toxic metabolites, and pH changes.
• Maintaining sterility ensures reproducibility and survival of cultures.

Key components of aseptic technique:

• Sterilization of media, vessels, and instruments (autoclaving).
• Surface sterilization of explants.
• Working within a laminar air flow (LAF) cabinet.
• Flame-sterilizing instruments between operations.
• Personal hygiene and clean lab practices.

An explant is an excised piece of living tissue or organ taken from a donor plant (mother plant) and used to initiate an in vitro culture.

Factors to consider during explant selection:

• Genotype of the plant: Some species/varieties respond better to culture (regeneration capacity).
• Age of the source tissue: Younger, actively dividing (meristematic) tissues have higher totipotency and regeneration ability.
• Physiological state: Actively growing plants in the appropriate season give better results.
• Type of organ/tissue: Shoot tips, nodal segments, leaves, roots, embryos, anthers, etc.
• Size of the explant: Larger explants survive better but carry more contamination risk; smaller ones are cleaner but harder to establish.
• Position on the plant: Apical regions are generally cleaner and more responsive.
• Health of the donor plant: Disease-free, well-nourished plants reduce contamination and improve success.
• Season/environmental conditions: Field-grown vs greenhouse-grown material affects contamination load.

Explant preparation involves a series of steps to obtain clean, viable tissue suitable for culture initiation.

Step-by-step process:

1. Selection of source material: Choose healthy, disease-free donor plants and appropriate tissue (shoot tip, node, leaf, etc.).
2. Pre-treatment/washing: Wash the tissue thoroughly under running tap water to remove dust, soil, and surface debris. Use a few drops of mild detergent (e.g., Tween-20) to remove waxy coatings.
3. Trimming: Cut the material to an appropriate size, removing damaged, senescent, or unnecessary parts.
4. Surface sterilization: Treat with disinfectants (e.g., ethanol, sodium hypochlorite, mercuric chloride) under aseptic conditions.
5. Rinsing: Rinse thoroughly with sterile distilled water (3–4 times) to remove residual sterilant which is toxic to tissue.
6. Final trimming: Under the laminar flow hood, cut away the damaged/sterilant-affected edges to expose healthy tissue.
7. Inoculation: Place the prepared explant onto the culture medium aseptically.

Note: All steps after surface sterilization are performed inside a laminar air flow cabinet to maintain sterility.

Surface sterilization is the process of removing or killing microorganisms present on the surface of an explant using chemical disinfectants without damaging the internal plant tissue.

Common chemical sterilants:

Key points:

• Ethanol is a rapid but harsh sterilant, often used as a brief pre-treatment.
• Sodium hypochlorite is the most widely used due to low toxicity to tissues.
• Mercuric chloride is highly effective but extremely toxic and non-biodegradable; requires careful disposal.
• A surfactant (e.g., Tween-20) is often added to improve contact by reducing surface tension.
• Thorough rinsing with sterile water afterward is essential to remove toxic residues.

Comparison of Sodium Hypochlorite vs Mercuric Chloride:

Conclusion:

• NaOCl is the sterilant of choice for most routine work owing to its safety and ease of use.
• HgCl₂ offers superior sterilization but its severe toxicity and environmental hazard restrict its use to difficult-to-sterilize explants only.

A Laminar Air Flow (LAF) cabinet is an enclosed workstation that provides a sterile, particle-free working environment for aseptic operations.

Construction:

• Cabinet/enclosure: Made of stainless steel with a transparent front shield.
• Pre-filter: Traps large dust particles from incoming air.
• Blower/fan: Draws and pushes air through filters.
• HEPA filter (High Efficiency Particulate Air): Removes 99.97% of particles ≥ 0.3 µm, including bacteria and fungal spores.
• UV germicidal lamp: Sterilizes the interior surface before use.
• Work surface: Flat platform for performing operations.

Working principle:

• Air is drawn in, passed through the pre-filter and then the HEPA filter.
• Sterile filtered air flows in a uniform, unidirectional (laminar) stream across the work area.
• This continuous positive airflow sweeps away airborne contaminants, preventing them from settling on the explant or media.

Types:

• Horizontal LAF: Air flows toward the operator (protects the sample).
• Vertical LAF: Air flows downward (protects both sample and operator).

Importance:

• Provides a contamination-free zone for cutting explants and transferring them to media.
• Reduces culture loss due to airborne microbes, ensuring high success rates in inoculation.

Aseptic inoculation is the transfer of a sterilized explant onto culture medium under sterile conditions.

Step-by-step procedure:

1. Preparation of LAF: Switch on the UV lamp for 15–20 min, then turn it off. Turn on the blower and allow airflow to stabilize (~15 min). Wipe the surface with 70% ethanol.
2. Arrange materials: Place sterilized instruments, media vessels, sterile water, spirit lamp/bead sterilizer, and Petri dishes inside the cabinet.
3. Hand sterilization: Wash hands and spray/wipe with 70% ethanol.
4. Instrument sterilization: Dip forceps and scalpel in ethanol and flame them; allow to cool before use to avoid tissue damage.
5. Explant trimming: Place the surface-sterilized explant in a sterile Petri dish and trim off the damaged/sterilant-affected ends using sterile forceps and scalpel.
6. Opening the vessel: Loosen the cap/plug of the culture tube; briefly flame the mouth of the vessel.
7. Transfer: Using sterile forceps, gently place the explant onto the medium with correct orientation (polarity). Ensure good contact with the medium.
8. Resealing: Flame the mouth of the vessel again and close it with the sterile cap/cotton plug.
9. Labeling: Label with date, explant type, and medium.
10. Incubation: Transfer cultures to the incubation/growth room under controlled light and temperature.

Precautions:

• Flame instruments between each explant to avoid cross-contamination.
• Avoid unnecessary movement of hands over open vessels.

Sterilization vs Disinfection:

In tissue culture:

• Sterilization is applied to nutrient media (autoclaving at 121°C, 15 psi, 15–20 min) and glassware.
• Disinfection (surface sterilization) is applied to the explant, since harsh sterilization would kill living plant tissue.

Endophytic contamination refers to contamination caused by microorganisms (bacteria/fungi) that reside inside the plant tissues, in intercellular spaces or vascular tissues, rather than on the surface.

Difference from surface contamination:

Control strategies:

• Use of antibiotics (e.g., cefotaxime, rifampicin) or antifungal agents in the medium.
• Selecting explants from actively growing meristematic tips (usually microbe-free).
• Indexing the mother plant for internal microbes before culture.
• Growing donor plants in controlled, clean greenhouse conditions.
• Applying thermotherapy or meristem culture to obtain clean tissue.
• Pre-treatment of donor plants with systemic bactericides/fungicides.

Surfactants (wetting agents) are substances added to sterilizing solutions to improve the efficiency of surface sterilization.

Common example: Tween-20 (polysorbate 20); also Teepol, Triton-X, or a few drops of liquid detergent.

Roles/functions:

• Reduce surface tension of the sterilant solution, allowing it to spread evenly over the explant surface.
• Improve wetting of hydrophobic (waxy/hairy) plant surfaces, ensuring the sterilant reaches all crevices.
• Enhance contact between the disinfectant and microorganisms, increasing sterilization effectiveness.
• Help remove air bubbles trapped on the surface (e.g., in trichomes/hairs) that shield microbes.
• Aid in the removal of surface debris and waxy coatings.

Usage:

• Typically 1–2 drops per 100 mL of sterilant solution.
• Explants may also be washed with a detergent solution before sterilization as a pre-treatment.

Precaution: Excess surfactant can be toxic to delicate tissues, so it must be used sparingly and rinsed off thoroughly.

After treating explants with chemical sterilants, rinsing with sterile distilled water (usually 3–4 times) is an essential step.

Reasons for rinsing:

• Removal of toxic residues: Sterilants like NaOCl and HgCl₂ are phytotoxic; residual chemicals can kill or damage the explant tissue.
• Prevents necrosis/browning: Retained sterilant causes tissue necrosis, bleaching, and death.
• Neutralizes chemical action: Stops the continued action of the sterilant so it does not over-treat the tissue.
• Improves survival and regeneration: Clean, chemical-free tissue has a higher chance of growth in vitro.

Important points:

• Water used must be sterile (autoclaved) to avoid re-contaminating the explant.
• Rinsing is done inside the laminar air flow cabinet under aseptic conditions.
• Especially critical for mercuric chloride, which is very toxic and requires multiple thorough rinses.

Autoclaving is a moist-heat sterilization method that uses saturated steam under pressure to kill all microorganisms including heat-resistant spores.

Standard conditions:

• Temperature:
• Pressure: (≈ )
• Time: minutes (longer for larger volumes)

Working principle:

• Water is boiled to generate steam inside a sealed chamber.
• As pressure rises, the boiling point of water increases to 121°C.
• The high-temperature steam denatures proteins and enzymes of microorganisms, killing them.

Procedure:

1. Fill autoclave with sufficient water.
2. Load media flasks/tubes (loosely capped) and wrapped glassware.
3. Seal the lid and allow steam to build up, expelling trapped air.
4. Maintain 121°C/15 psi for the required time.
5. Allow slow cooling and pressure release before opening.

Items sterilized: Nutrient media, distilled water, glassware, forceps, scalpels, cotton plugs.

Note: Heat-sensitive substances (some vitamins, hormones, antibiotics) are sterilized separately by membrane filtration (0.22 µm filter) and added to cooled media.

Maintaining asepsis requires strict adherence to good laboratory practices at every stage.

Personal precautions:

• Wear clean lab coats, gloves, and masks.
• Wash and sterilize hands with 70% ethanol before working.
• Avoid talking, coughing, or sneezing over open cultures.

Work area precautions:

• Perform all transfers inside the laminar air flow cabinet.
• UV-sterilize the cabinet before use and wipe with 70% ethanol.
• Keep the transfer area clean and free from unnecessary items.

Instrument precautions:

• Flame-sterilize forceps and scalpels between operations; cool before use.
• Use a bead sterilizer or ethanol dip for repeated sterilization.

Media & vessel precautions:

• Properly autoclave all media and glassware.
• Flame the mouths of culture vessels while opening and closing.
• Use sterile distilled water for rinsing.

General lab precautions:

• Maintain a dust-free, air-conditioned environment.
• Regularly clean floors and surfaces with disinfectant.
• Restrict entry and fumigate the transfer room periodically.
• Dispose of contaminated cultures promptly by autoclaving.

Explant polarity refers to the inherent axial orientation of plant tissues, having a distinct apical (shoot/top) end and a basal (root/bottom) end, based on the natural direction of growth and hormone (auxin) transport.

Basis of polarity:

• Plant cells exhibit polar auxin transport (from apex to base).
• This creates a physiological gradient that determines where shoots and roots form.

Importance during inoculation:

• Correct orientation is essential for proper regeneration.
• The basal end should generally be inserted into or in contact with the medium so roots develop downward and shoots grow upward.
• Inverted (upside-down) placement can lead to poor growth, delayed regeneration, or abnormal development.
• Ensures proper nutrient and hormone uptake from the medium.

Practical note:

• For stem/nodal segments, the morphological orientation must be maintained.
• For some explants (e.g., leaf discs), polarity is less critical but the adaxial/abaxial surface contact may affect response.
• Marking the apical end during excision helps maintain correct polarity.

Contamination is the major problem in tissue culture. It can arise from several sources.

Sources of contamination & their control:

1. The Explant (surface & internal):

• Source: Microbes on/in donor plant tissue.
• Control: Effective surface sterilization, use of meristem tips, antibiotics, disease-free mother plants.

2. Culture Media:

• Source: Improperly sterilized or contaminated media.
• Control: Proper autoclaving (121°C, 15 psi, 15–20 min); filter-sterilize heat-labile additives.

3. Glassware & Instruments:

• Source: Unsterile tools and vessels.
• Control: Autoclave/flame-sterilize; use bead sterilizer.

4. Air (airborne spores):

• Source: Dust and microbes in laboratory air.
• Control: Work in HEPA-filtered LAF cabinet; fumigate and UV-treat the room.

5. The Operator (human handling):

• Source: Hands, breath, hair, clothing.
• Control: Sterilize hands with ethanol, wear lab coat/mask/gloves, avoid talking over cultures.

6. Mites and Insects:

• Source: Carry fungal spores into cultures.
• Control: Maintain clean, pest-free culture rooms; seal vessels properly.

Conclusion: A combination of proper sterilization, aseptic technique, and clean facilities is required to minimize contamination.

Antibiotics and antifungal agents are sometimes incorporated into the culture medium to control persistent bacterial and fungal contamination, especially endophytic microbes.

Commonly used antibiotics:

• Cefotaxime, Carbenicillin, Rifampicin, Streptomycin, Ampicillin – used against bacteria.

Common antifungal agents:

• Nystatin, Amphotericin B, Benomyl, Bavistin (carbendazim) – used against fungi.

Mode of application:

• Added to the medium (usually filter-sterilized as they are heat-labile).
• Or used as a pre-treatment dip for explants.

Limitations:

• Phytotoxicity: Many antibiotics are toxic to plant cells and can inhibit growth/regeneration.
• Selective action: Effective only against specific microbes; broad-spectrum control is difficult.
• Development of resistance: Prolonged use can select for resistant strains.
• Cost: Some antibiotics are expensive.
• Not curative for all: Cannot fully eliminate deep-seated endophytes.
• Heat-sensitivity: Cannot be autoclaved; require separate sterile filtration.

Conclusion: Antibiotics are a supplementary control measure and cannot replace proper aseptic technique and surface sterilization.

Both techniques exploit the fact that the apical meristem region is generally free of viruses and microbes.

Basis of virus-free status:

• The apical meristem has no fully developed vascular tissue, and viruses move mainly through the phloem, so the actively dividing tip cells escape infection.

Conclusion:

• Meristem culture is chosen for producing disease-free clean stock, while shoot-tip culture is favored for mass multiplication with easier handling.

Pre-treatment/washing is a preparatory step performed before chemical surface sterilization to reduce the contamination load and enhance sterilization efficiency.

Significance:

• Removes surface debris: Soil, dust, and particulate matter carrying microbes are washed off.
• Reduces microbial load: Fewer surface microbes mean better sterilization outcomes.
• Removes waxy/hydrophobic coatings: Improves penetration of the sterilant.
• Loosens firmly attached contaminants in trichomes and crevices.
• Improves overall success rate of aseptic establishment.

Agents/methods used:

• Running tap water: Prolonged washing (30 min – few hours) to remove debris.
• Mild detergent / liquid soap (e.g., a few drops of Tween-20 or Teepol): removes wax and grease.
• Soft brush: to physically scrub hairy or rough surfaces.
• Fungicide/bactericide dip (e.g., Bavistin solution): for heavily contaminated field-grown material.
• Ethanol wipe (70%): brief pre-disinfection of the surface.

Note: After washing, explants are trimmed and then subjected to the main surface sterilization procedure.

Contamination in tissue cultures is usually detected by visual observation within a few days of inoculation.

Symptoms of Bacterial Contamination:

• Appears as slimy, wet, glistening, cream/white/yellow colonies or films.
• Often spreads over the medium surface and around the explant.
• May produce a cloudy/turbid appearance in liquid or on solid media.
• Sometimes accompanied by a foul odor.
• Usually appears rapidly (within 1–3 days).

Symptoms of Fungal Contamination:

• Appears as fluffy, cottony, or filamentous (mycelial) growth.
• Colored spores/mycelia — white, black, green, grey, or pink.
• Grows in circular, spreading patches, often aerial.
• May appear slightly later than bacterial contamination.

Visual distinction:

Action: Contaminated cultures should be removed immediately and autoclaved before disposal to prevent spread.

Complete Protocol for Shoot-tip Culture Establishment:

A. Explant Selection & Preparation:

1. Select young, healthy, actively growing shoot tips from a disease-free field-grown plant.
2. Excise shoots ~2–3 cm; remove large leaves.
3. Wash under running tap water for 30–60 min to remove soil/debris.
4. Immerse in detergent solution (a few drops of Tween-20) for 5–10 min; rinse well.
5. Optional: dip in fungicide (0.1% Bavistin) for 15–30 min to reduce fungal load; rinse.

B. Surface Sterilization (perform inside LAF):

6. Treat with 70% ethanol for 30–60 sec.
7. Rinse once with sterile distilled water.
8. Immerse in 0.1% mercuric chloride for 3–5 min OR 1–2% sodium hypochlorite for 10–15 min (add 1–2 drops Tween-20).
9. Rinse 3–4 times with sterile distilled water to remove all sterilant residues.

C. Aseptic Inoculation (inside LAF cabinet):

10. Sterilize LAF with UV (15–20 min) and wipe with 70% ethanol; sterilize hands.
11. Place sterilized shoot tip in a sterile Petri dish; under a dissecting microscope, trim to expose the healthy shoot apex.
12. Flame-sterilize forceps/scalpel; cool before use.
13. Flame the mouth of the culture tube; place the explant on the medium (e.g., MS medium + BAP) with correct polarity.
14. Flame the mouth again and close with sterile cap.
15. Label with date, explant, and medium details.

D. Incubation:

16. Incubate at 25 ± 2°C under 16 h light / 8 h dark photoperiod.
17. Observe daily for contamination and record growth response.

Precautions: Flame instruments between explants; maintain strict asepsis throughout; discard and autoclave any contaminated cultures.