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Freeze & Thaw lysis with XNA

1. spin down bacteria, take 5uL of pellet, add to 20uL of XNA Extraction Solution
2. Freeze in -20 for 5 min
3. Thaw for a minute, room temperature or in your fingers
4. repeat freeze – thaw two more times
5. add XNA Dilution solution

General bacterial cell lysis

http://www.accessexcellence.org/LC/SS/PS/PCR/PCR_technology.php:

1. For bacterial samples take a toothpick and scrape the teeth, or swab the throat, ears or between the toes. Resuspend material in 500ul of water. Freeze and thaw sample three times with vigorous shaking or vortexing between repetitions to break the bacterial cell wall. Although not all DNA will be released from the cells, there will be a sufficient quantity for PCR.
2. Place the sample in a 95oC heating block, or in boiling water, for 5 minutes. This step inactivates the DNase molecules that are found in the sample preparation. If left intact, DNase could clip the desired DNA template molecule into fragments which would be unsuitable for PCR. If there is very little DNA in the sample preparation, the DNA can be concentrated by ethanol precipitation. The sample is now ready for PCR

Actinomycete DNA Extractions

1. Mycelium was collected from plates and suspended in 500 μL of cetyltrimethylammonium bromide buffer (Hillis et al. 1996) in microcentrifuge tubes.
2. Cells were broken by cycles of freezing and thawing (they were dipped in liquid nitrogen and then placed in a heat block at 65 °C); this procedure was repeated at least 3 times.
3. One volume of chloroform was added, tubes were vortexed briefly, and then centrifuged for 10 min at 10 000g.
4. Precipitation with 1 volume of 100% isopropanol was carried out overnight at –20 °C.
5. After centrifugation, the DNA pellet was washed twice with 70% ethanol and air dried. DNA was resuspended in 50 μL of buffer TE/10 (10 mmol Tris–HCl/L, 0.1 mmol EDTA/L; pH 8.1); aliquots were diluted (1:200) in ddH2O and used for PCR amplification (Cafaro & Currie, 2005).

Protocol

Dilution to Extinction
Increases culture time, decreases inter-colony interactions, increases observed diversity over continuous-surface culturing (Collado et al., 2007). Great for common but fastidious, slow-growing organisms.

Protocol

Cellulose Degradation
Isolate possible cellulose degraders on CMC (5 g/L & salts, false positives possible) or MCC (4g/L & salts). Use CMC media for Congo Red test – grow isolate in the plate center for a week, overlay with Congo Red, wash with water, wash with salts (3M NaCl), check for clearing. When isolating cellulose degrading bacteria from wood, it’s good to add antifungal antibiotics, otherwise Trichoderma takes over.

Preparing media

1. after the plates have cooled, mark each stack with a line of the appropriate color (easy when plates are stacked)
2. put them back in sleeves and tape those over
3. write the media type, your name, and date on the tape
4. clean up the hood completely! The lab that owns the hood has started to use it frequently again, so we need to make sure that we always leave it immaculate.
5. store the new plates in the fridge in the teaching storage.

TSA media

Tryptic Soy Agar (same as Trypticase Soy Agar) (red stripe on plates)

• 40g of BD TSA powder
• 1L water of purified water

Autoclave, pour plates, stripe with red marker.

Nutrient Agar

(blue stripe on plates):

• 23g of Nutrient Agar powder
• 1L of purified water

Autoclave, pour plates, stripe with blue marker.

The following section refers only to media for culturing filamentous actinomycetes

Actinomycetes like dry media – let plates dry out before plating. Keep media with antibiotics in a cold and DARK place, to prevent degradation.

Cafaro & Currie 2005: Inoculum from an insect transferred onto chitin-agar plates, containing nystatin (10,000 units/mL). After 3–5 weeks of growth at room temperature, bacterial colonies are subcultured onto Czapek yeast autolysate agar with antibiotics (nystatin 10 000 units/mL and cycloheximide 5% w/v) (Cafaro & Currie, 2005).

An isolate is cultured in YMEA (4 g yeast extract, 10 g malt extract, and 4 g glucose per 1 L) at 30˚ C for 3 days. 20 mL of the YMEA culture was then inoculated to 200 mL of YPM (2 g yeast extract, 2 g peptone, and 4 g mannitol per 1 L). The YPM culture was incubated for about 15 h (for a subsequent use in spectroscopy) (Scott et al., 2008).

Humic acid-vitamin agar for actinomycete endophytic actinomycetes amended with nalidixic acid to suppress other bacteria (Hayakawa & Nonomura, 1989) and by cycloheximide and nystatin against fungi (Taechowisan et al., 2003).

(Crawford et al., 1993) tested many types of media for isolation of root endophytic actinomycetes, and found that the best was the nutrient-poor Water-Yeast-Extract agar (WYE) modified from that of Reddi and Rao) contained yeast extract (Oxoid; 0.25 g/liter) as the sole carbon and nitrogen source and agar (Oxoid; 18.0 g/liter). The medium was buffered with K2HPO4 (0.5 g/liter). (Chen et al., 2009) used the same media to isolate Pseudonocardia spp. from wood, used tap water (“TWYE” agar) and containing nalidixic acid (10 mg/ml), nystatin and cycloheximide (each at 50 mg/ml).

WYE does work for actinomycetes, but it tends to remain more damp than chitin media, so they germinate with lower frequency.

100 μg nystatin and cycloheximide/ml (Taechowisan et al., 2003)

Nystatin can be probably substituted with Natamycin (=Pimaricin). Soluble in DMSO. Sigma-Aldrich suggests not to autoclave or filter Nystatin solution.

PBS buffer can be substituted with 0.1 % Tween (which one?)

Emerson Media

(red & orange stripe on plates):

1. Mix all ingredients except starch.
2. Microwave starch in ~50 ml dd H2O until transparent (~40 seconds).
3. Pour dissolved starch into mixture.
4. Autoclave.

Equipment for field sampling

• autoclaved screw-top vials with moist paper (if sampling live beetles)
• autoclaved empty screw-top vials
• ethanol
• squirt bottle (or separate vials with ethanol)
• bullet box for vials
• larger tubes
• chainsaw + gas + oil
• 2 broad chisels
• crow bar
• anvil pruner
• folding saw
• hammer
• hatchet
• soft forceps on a string
• hard forceps
• scalpel
• scalpel blades
• lighter (for forceps & scalpel sterilization)
• ziploc bags
• permanent markers (alcohol-resistant)
• pigma pen for labels
• notebook

In the field

1. beetles: pierce head, put in numbered vial, record
2. larvae & frass: put in numbered vial or tube, record
3. fungi: same as above
4. frontalis: small samples tied up with rubber band, in ziploc bag

Sampling bacteria from beetles

Up to 3 larval chambers, and up to 3 adult beetles (ideally new generation) per tree

1. surface wash 2. the beetle crushed (elytra removed, surface-washed in bleach, water, ethanol, and dried out)

Materials for dissection

(all autoclaved)

• eppendorf vials
• pestles
• (chisel)
• pruner
• pipette
• 1000 uL pipette tips
• EtOH bath
• scalpel
• vial rack
• surface wash for adult beetles (elytra removed, surface-washed in bleach, water, ethanol, and dried out)
• buffer