Fact Sheet Fusion

Picture format

• tiff with LZV compression or a high quality jpg.
• 750px wide, height is unimportant but a consistent height within a taxon is an aesthetic benefit
• specimens directed to the left
• white background, if possible
• both dorsal and lateral aspects, plus one or two diagnostic characters

Fixating

For visualizing mandibular mycangia, beetle heads or bodies should be fixed in 96% ethanol, immersed in 30% hydrogen peroxide for 24 h, and embedded in paraffin.

Microtome

Preparates can be sectioned on a microtome at 5 μm and stained with hematoxylin (to emphasize nuclei in fungi) and eosin (to visualize proteins, muscles, and other beetle tissue).

Important points

1. The immersion in hydrogen peroxide is critical for softening beetle exoskeleton and to avoid fracturing.
2. The angle of beetle immersion in the wax is also critical. It depends on what mycangia you are interested in, what angle you want to slice them, and what is the angle of attachment of the wax sample to your microtome. You may need to play with the beetle position as the wax is hardening.

Before introducing beetles into new tubes, pass them through several vials with moist kimwipe (all sterile), two days in each vial. This way they empty out their guts, attached spores, and mites. IMPORTANT – poke a hole in the vial lid, otherwise beetles quickly suffocate. No need to rush – the longer the beetles are out of the gallery, the more pure will be their symbiont in the mycangium.

Washing beetles

Prepare:

• vial with 1ml PBS + 0.1% Tween (one small drop), and
• vial with 40% EtOH (100% ethanol is too much, even 70%).

1. Drop beetle in PBS, vortex for 15 sec.
2. Take beetle out, put on paper to dry out.
3. Drop beetle in EtOH, keep in for 5 sec.
4. Take beetle out, put on paper to dry out.
5. If beetle is happy and alive, put in tube.
6. Record tube number and beetle species.

Protocol 1

Developed for X. crassiusculus in North Carolina.
Used sweetgum (Liquidambar). Cut out branches ~4cm thick. Autoclave, OR soak in ethanol and then autoclave without drying.
Direct inoculation:
1. Cut end of vial and plug with cotton (control for excessive moisture).
2. Drill shallow holes of equal diameter as vial thread.
3. Screw vial in, with beetle inside.
4. Place log in jar of water, parafilm around.
5. Place in well-lit and well ventilated space.

Indirect inoculation:
1. Parafilm log ends.
2. Drill hole in base of diameter equal to small aquarium tube.
3. Plug tube with cotton and connect tube to water source.
4. Insert in log.
5. Place on a moist paper, in a large screen enclosure.
6. Release beetles in enclosure.

Protocol 2

Cut fresh branch (50 x 5 cm) of sweetgum or maple, seal its exposed ends with parafilm (or wax, or natural latex) to prevent dessication. Prepare containment vials – between 1-3 cm, bottom perforated with many miniature holes, or with one large hole sealed over with wire micromesh (for ventilation – water condensation traps and kills beetles). Soak log for 24 hours in water, remove from water na dry briefly. Drill shallow holes of exactly the diameter of your vial. Place beetle in hole, cover with vial. Keep in humid place. Tupperware with source of water is sufficient. Ventilation is ideal, especially in the beginning of gallery development, but not essential.

For rearing beetle families in twigs: PDC broth (Potato dextrose + 10g casein/L). Tested, but results unclear. Some beetles emerged, not many.

1. For one beetle, assemble thirteen 1.5 mL micro centrifuge tubes containing .5 mL of sterile PBS and label them as following:

(W)
(SW.1),  (SW.01), (SW.001)
(MY.1),  (MY.01), (MY.001)  OR   (G.1),  (G.01), (G.001
(HP.1),  (HP.01), (HP.001)
(A.1),  (A.01), (A.001)

W=Water, SW=Surface Wash, MY=Mycangium, G=Gallery, HP=Head+ pronotum, A=Meso+metathorax+abdomen

1. Add one drop of tween oil only to (SW.1).
2. Wipe down microscope area with ethanol. Put the following items under UV sterilization for at least 10 minutes:
   1. Weak forceps, strong forceps, paraffin, minuten pins, 000 pins, and centrifuge tubes (see below). Open the cap of the tube with tween oil.
3. Add the beetle to the tube with tween oil (SW.1). Vortex for 20 seconds at 2100 rpm.
4. Remove the beetle from the tube with tween and add to a tube with sterile water or PBS (W). Vortex for 20 seconds at 2100 rpm.
5. Transfer the beetle with sterile weak forceps to a dry kimwipe, and then onto the paraffin.
6. Secure beetle with dorsal side up using minuten pins. Place four pins total in between the mesonotum and pronotum. Two of these pins should almost form a \/ shape in the parafilm, making a cradle for the beetle. The other two pins should form a /\ shape, holding the beetle down and preventing it from rotating.
7. Insert a minuten pin into the anterior portion of the pronotum. Once inserted, decrease the angle the pin so that it is parallel to the paraffin and so that the pronotum of the beetle slides forward, revealing the mycangium. Secure this pin in the pronotum down with more pins to maintain its position.
8. Using a sterile 000 pin, work the edge of the mycangial membrane to lift up the entire mycangium as a cohesive mass. The color is typically orangish or beige in compactus, and the texture looks like very fine sand or eggs. Transfer the entire mycangium into a centrifuge tube. This may take a few transfers.
9. Using sterile scalpel, sever the beetle just anterior to the location of the mycangium. Transfer the head and pronotum into one tube, and the rest of the beetle into another tube.

Materials

• grinding tool
• scalpel
• hood time
• strong forceps
• vials
• ethanol in petri dish
• vial rack

Procedure

Scrape piece of 1 gallery in the hood, scrape into

a. 1 ml of 1X PBS AUTOCLAVED phosphate buffer saline solution in tube

3. mortar & pestle
4. plate directly, refrigeration decreases growing success

5. Plate three different dilutions, refer to the appropriate protocol.