Freezing

• glycerol (most fungi an bacteria do well in 10%, some need 20%) or mineral oil (messy)
• Put three agar plugs (or just chunks of mycelium, or one big chunk, whatever is easier) from each plate into its respective storage vial. Agar plugs (cubes with fungus) or pieces of mycelium can be cut with scalpel; make sure that scalpel is perfectly sterilized after each culture. Take the growing edge of the fungus, or a whole growing colony, not the old crusty center.
• If you are using minislants – just pour the sterile 20% glycerol in the minislant tube
• Place in Mr. Frosty (blue/white container on top of the fridge). Fill up the bottom compartment with isopropyl alcohol (in the chemicals storage). Put in -80C freezer. It will freeze slowly, 1C per minute.
• Record in the Scolytos Database!!! Each isolate will get genus name “fungus” and its unique Species name will be its Isolate name from the isolations database. In “count”, record 1.

Reviving

Prepare:

• vials with 1mL PBS. (Label them with numbers corresponding to frozen samples)
• equal number of PDA plates (label these with regular database numbers)
• sterilizer, scalpel

1. Take out sample. 2. Cut gel disk inside the tube with carefuly sterilized scalpel. 3. Use only one half (other half stays). 4. Put half-disk in vial with PBS, shake briefly to rinse off surplus glycerol or mineral oil, and put on PDA plate.

Using multimode detector (plate reader) Beckman Coulter DTX 880 to measure absorbance at 630nm with a 20×20 detector grid, using sterile 24-well Costar plate with a lid against contamination, 1 ml of media per well. Measured on the first day and on fourth day, comparing (cellulose media minus negative control) versus (positive control minus negative control).

Vortex chunk of fungus mycelium, take 0.1 ml of supernatant, add to 0.9 ml of media.

Assays

Filter paper assay

Testing for utilization of pure unaltered cellulose

• control-noble (neg. control)
• filter paper
• PDA (pos. control)

CMC & Congo Red assay

Testing for cellulose-degrading enzymes

• control-salts (neg. control)
• CMC-salts
• CMC-PDB_0.25
• dextrose-salts
• PDA (pos. control)

MCC assay

Testing for cellulose carbon utilization (i.e., growth)

• control-noble (neg. control)
• MCC-salts
• MCC-PDB_0.25
• PDA (pos. control)

Media needed (all media per 1L, all with 20ml/L Penicillin&Streptomycin):

• control-noble: 15g noble agar, salts* 0NS
• control-salts: 15g normal agar, salts* 0S
• MCC-salts: 5g MCC, 15g noble agar, salts* MNS
• MCC-PDB_0.25: 5g MCC, 0.25g PDB, 15g regular agar M-green
• CMC-salts: 5g CMC, 15g normal agar, salts* CS
• dextrose-salts: 5g dextrose, 15g normal agar, salts*
• CMC-PDB_0.25: 5g CMC, 0.25g PDB, 15g regular agar C-green
• filter paper: piece of autoclaved Whatman filter paper 15g regular agar, salts* 0S

salts*

(prepare 10x solution ahead= 10X of the following per 1L, autoclave):

Salts	Amount
KH2PO4	0.2 g
NH4Cl	0.25 g
KCl –	0.5 g
CaCl2	0.15 g
NaCl	1 g
MgCl	0.6 g (or 1.2g if hydrated)
K2SO4	2.84 g

include in media through filter after autoclaving: 1ml of 1000x trace minerals solution 10ml of 100x trace vitamins solution

Media based on Czapek-Dox

Czapek-Dox-CMC (cellulose degradation test)

Czapek-Dox neg. control (no carbon source)

Czapek-Dox-dextrose (positive control) Czapek-Dox neg. control (no carbon source)

The Czapek-Dox doesn’t work for Raffaelea – doesn’t grow on positive control. Try different media – Czapek-Dox with proteins, or with little bit of yeast extract, or dextrose+salts+NaNO3.

Yeast Nitrogen Base media – Liquid

Tested media:

1) Prepare YNB&AA& antibacterials 10x stock: 6.7g YNB&AA and 20mL Streptomycin/Penicillin concentrate, in 100mL water. Use warm water, but do NOT autoclave! Store in dark and cold place.

2)YNB- CMC-liq (do not use MCC – not soluble, not usable in absorbance measurements)

• water 90mL
• CMC 3g
• after autoclaving: 10ml YNB&AA&antibacterials 10x stock through filter. pH: 5.7

YNB- dex-glu-liq

• water 90ml
• dextrose 1.5g
• glucose 1.5g
• after autoclaving: 10ml YNB&AA&antibacterials 10x stock through filter

YNB- blank-liq

• water 90mL
• after autoclaving: 10ml YNB&AA&antibacterials 10x stock through filter

Yeast Nitrogen Base media

1) prepare YNB&AA 20x stock: 26.8 g in 200mL warm water, do NOT autoclave! Store in dark and cold place.

2) autoclave 50 pieces of filter paper (the same size as last time) as you are autoclaving the media

3) media:

YNB-CMC

• water 950 mL
• agar 15g
• CMC 30g
• after autoclaving: 50ml YNB&AA 20x stock through filter

YNB-dex-glu

• water 950ml
• agar 15g
• dextrose 15g
• glucose 15g
• after autoclaving: 50ml YNB&AA 20x stock through filter

YNB-filter_paper

water 950 mL

• agar 15g
• one piece of filter paper,
• after autoclaving: 50ml YNB&AA 20x stock through filter

YNB-blank

• water 950mL
• agar 15g
• after autoclaving: 50ml YNB&AA 20x stock through filter

Congo Red assay

Prepare Congo Red solution (1g/L) and 1M solution of NaCl (58g/L). Overlay plate with Congo Red for 15 minutes, pour off, overlay with NaCl for 15 minutes, pour off, look for zone of clearing

Protocol

Materials

• 1000ul and 100ul pipettes
• pipettes tips
• two pairs of fine hard forceps
• petri dishes for beetle dissection
• microscope in hood
• vials
• vial rack
• sterile 1X PBS
• alcohol-resistant marker

General notes

1. Wash the beetle with a surfactant and/or saline, to get most of the dirt off. You don’t need to get every last fungal spore off the surface (we will dilute those away). You also don’t need to use any toxic solutions, often used in older works. Some people use ethanol for surface sterilization; it probably helps in removing some contaminants, but there is also a high percentage of the mycangial symbiots that die. We have tested it…
2. Don’t grind up the whole beetle – focus on the right body part. If the mycangium is in the head, use the head (most Xyleborini). If it’s in the prosternum (for example, Xyloterini, Corthylini) or pronotum (Platypodinae), use the prothorax. If it’s in the mesonotum (the Xylosandrus–Anisandrus clade of Xyleborini) it gets a bit trickier, but with a little practice you will learn how to excise that general part of the body out as well. The main point is to avoid most of the surface, the alimentary canal (particularly the hind gut which is full of yeasts) and the space under elytra, which also hosts many unwanted associates (including nematodes). Yes – the space under the elytra is dirty. When you want to study the microbial associates of people’s mouth, you also don’t grind up the whole person.
3. Dilution plating! This is the ESSENTIAL part of the process. Plate several dilutions of your inoculum, and use the lowest one where some Colony Forming Units end up growing in the end. The goal here is to dilute away all the non-specific associates which are likely present in lower abundances, and only capture the abundant symbionts. Those are typically present in thousands of cells, so you have a good chance of getting mostly those in the lowest dilution.

Dilution plating

1. Prepare two tubes per each sample, label them “0.1” and “0.01”, fill each with 500 ul of water or PBS.
2. Suspend mycangium in the tube “0.1” and vortex.
3. Plate 50 ul of the suspension on media. Record the plate as “0.1 dilution” in [PLATES].[note] in the Isolations database.
4. Transfer 50 ul of the initial suspension to the second tube (“0.01”) and vortex.
5. Plate 50 ul of the second suspension on second media plate, and record that plate as “0.01 dilution”.
6. Plate 5 ul of the second suspension on a third plate, and record it as “0.001 dilution”.

Quantititive extraction for fungus community characterization:
See Calculating colony-forming units