Introduction: Reading Agar Plates
Agar work is a visual skill. Before you can transfer, select sectors, or identify contamination, you need to be able to read what is happening on a petri dish. This guide walks through each stage of agar work with detailed descriptions of what you should be observing and what actions to take. Consistent observation habits — checking plates at the same time each day and noting changes — build the pattern recognition that makes agar work intuitive over time.
The most important habit in agar work is to observe plates from multiple angles and lighting conditions. Viewing a plate from the bottom (through the agar) reveals the depth of mycelium penetration and makes early contamination visible as colour changes in the gel. Viewing from above at a low angle in strong side-lighting reveals surface texture — the difference between rhizomorphic and tomentose growth becomes immediately apparent in raking light.
Part 1: Agar Media — Recipes and Selection
1 Malt Extract Agar (MEA) — The Standard
MEA is the most widely used agar media for mushroom cultivation. The malt extract provides sugars and trace minerals that support healthy mycelium growth, while agar provides the solid matrix. The amber colour of MEA makes contamination — particularly bacteria and Trichoderma — visually obvious against the background.
Standard MEA recipe: 10–20 g malt extract powder + 20 g agar powder per 1 litre of water. Mix cold, pour into media bottles, pressure cook at 15 PSI for 20–30 minutes, cool to 50°C, pour plates.
Higher malt concentrations (20 g/L) produce faster mycelium growth but can make contamination harder to spot and give Trichoderma a competitive advantage. Starting at 10 g/L is safer for new cultivators.
2 Other Common Agar Recipes
| Media | Recipe (per litre) | Best For |
|---|---|---|
| MEA | 10–20 g malt extract, 20 g agar | General isolation; most species |
| PDA | 200 g potato (boiled), 20 g dextrose, 20 g agar | Faster growth; good for vigour assessment |
| LME (Light MEA) | 5 g malt extract, 20 g agar | Slower growth; easier to spot early contamination |
| Oatmeal agar | 30 g oats (boiled), 20 g agar | Slightly contam-resistant; good for Pan Cyan |
Part 2: Pouring Plates — Technique and Common Errors
1 Cooling the Media Correctly
After pressure cooking, allow the agar media bottle to cool until it is comfortably warm to the touch — approximately 45–50°C (113–122°F). At this temperature the agar is still fully liquid but will not generate the massive condensation that hot agar produces when poured into a cooler petri dish.
Too hot (above 60°C): Poured plates develop heavy condensation on the inside of the lid. This condensation drips back onto the agar surface and creates bacterial wet spots within 24–48 hours. You can flip plates and leave them upside-down during solidification to reduce this, but prevention through correct pour temperature is better.
Too cool (below 40°C): The agar begins to solidify in the bottle, producing lumpy plates with an uneven surface. Reheat the bottle in a warm water bath (not microwave) to return to liquid state if this happens.
2 The Stack Pour Technique
Work inside a still air box (SAB) with alcohol-wiped surfaces. Stack 10–20 sterile petri dishes. To pour the bottom dish, lift the entire stack with one hand — the stack itself shields the open dish from airborne contamination while you pour with the other hand. Pour approximately 15–20 ml per standard 90 mm dish. Immediately set the stack back down, trapping the poured lid back in position.
Pour plates in a single smooth session without pause — once a media bottle is open in the SAB, any hesitation gives airborne particles more time to settle. A typical session of 20–25 plates takes 8–12 minutes.
Part 3: Making a Transfer — Step-by-Step
1 Flame Sterilisation
Flame-sterilise the scalpel blade outside the SAB or away from alcohol fumes — the flame can ignite isopropyl alcohol vapour. Heat the blade until it is visibly red-hot (3–5 seconds in the flame). Bring the hot blade into the SAB and allow 5–10 seconds to cool by touching it briefly to an uninoculated area of agar. You should hear a brief hiss as the blade contacts the agar and immediately steams. This confirms the blade was hot enough to sterilise and has now cooled enough not to kill the mycelium.
2 Identifying and Selecting the Transfer Site
Examine the plate at a low angle under strong light. The leading edge of mycelium growth — the outermost expanding frontier — shows the most distinct sector variation. Rhizomorphic sectors appear as thin, rope-like or web-like strands extending further ahead than adjacent tomentose (fluffy) sectors.
Your target is a wedge cut from the very leading tip of the fastest-growing rhizomorphic sector — specifically the point approximately 3–5 mm behind the absolute growing edge. This zone contains the most recently generated, actively growing mycelium. Older, inner growth is less metabolically active and transfers less reliably.
Cut a wedge approximately 2–4 mm across — roughly the size of a grain of rice. The cut should go all the way through the agar to ensure you capture both surface mycelium and any mycelium that has penetrated into the agar gel.
3 Executing the Transfer
Open the destination plate as briefly as possible. Place the agar wedge face-down — mycelium side touching the fresh agar surface — at the exact centre of the new plate. Pressing the wedge into contact with the agar ensures the mycelium can begin growing immediately without needing to bridge an air gap.
Close the plate immediately and seal with Parafilm or micropore tape. Label with the source plate number, transfer generation (G1, G2, G3...), and date. Incubate at 23–26°C in a dark location. First growth should be visible within 2–4 days.
Part 4: Rescue Transfers and Saving Contaminated Plates
1 Assessing Whether a Rescue is Viable
A plate with contamination can sometimes be rescued if the contamination is visually distinct and has not reached the mycelium. The three criteria for a viable rescue are: (1) the contamination has not yet sporulated (no green, blue, or black powdery patches); (2) there is a clear gap of uncontaminated agar between the contamination and the cleanest mycelium; and (3) the contamination is in a localised zone, not distributed across the plate.
If Trichoderma has already turned green, do not attempt a rescue — the risk of spreading spores during the transfer outweighs any potential benefit. Seal without opening and dispose.
2 Executing a Rescue Transfer
Work with extreme care. The goal is to cut a wedge from the healthiest, cleanest mycelium — typically on the opposite side of the plate from the contamination — without disturbing the contaminated zone at all. Move slowly. Do not pass the scalpel over the contamination area on the way to your target.
Transfer to a fresh plate. You may need to perform 2–3 serial rescue transfers — each time selecting the cleanest growth from the new plate — before the culture is fully clear of contamination. Even after apparent success, monitor new plates closely for the first 7–10 days for any recurrence.