1. The Gut Guide: A handbook for the trichomycete morphology lab
Emma R. Wilson
Boise State University
Research lab of Merlin White, PhD

2. Contents Introduction Part 1: Overview of Trichomycetes
Symbiosis
Taxonomy
Part 2: Collecting Specimens
Where are they?
Aquatic collections
Sorting Hosts
Part 3: Maintenance, Dissection & Preservation
Keeping hosts alive
Dissection
Host preservation
Fungal preservation
Part 4: Culturing
Media recipes
Making media
Pouring Plates
Making Slants
Transferring cultures
Part 5: Morphometrics
Where and what to measure
Examples and tips
Conclusions
Appendices
1. Species list by Order
2. Host keys
3. Further reading

3. Introduction
This handbook is made for any student who would like to deepen their understanding of the curious world of trichomycete fungi and protists*.
We will start out by briefly introducing the biology of trichomycetes.
Next, we’ll give some examples and tips for specimen collection and how to be successful in the field.
After that we’ll focus on the tricho hosts and what to do with them once you’ve gathered them in the field.
One important aspect of trichomycetology is culturing. That will be covered in Part 4 of the guide.
Finally, we’ll go over morphometrics (a way to measure your tricho and determine what species you have).
At the end of this guide, you’ll find several appendices including a list of genera, host keys, and a link for further reading.
Enjoy!
* “trichomycete fungi and protists” will often be shortened to “trichos” (pr: “trick-Ohs”) throughout this handbook.

4. Web Resources
In addition to this guide, the following links are vital for anyone interested in trichos:
Created by Dr. Robert Lichtwardt – the modern father of trichos.
Lets go through each section to show some highlights of how best to use each page as a burgeoning trichomycetologist.
Bob – modern father of trichos? Guru of trichos? How to show is importance?

5. Web Resources
On the home screen of , there is a brief summary about trichos as well as two very useful links – the Monograph and the Lucid Keys – that are used by scientists in the lab nearly every day.
We’ll go into detail about these next
Shortcut to Lucid Keys
Shortcut to Monograph

6. Monograph
Contents are a great place to start navigating
Glossary contains many tricho-specific definitions
This page is essentially a book (originally published in 1986) whose current version is published on the web.
It contains a wealth of information from host specificity to molecular techniques with trichos
The easiest way to navigate the monograph is to click on “Contents” and choose from the listing on the page that appears.
There is also a great glossary for many tricho-specific terms.

7. Lucid Keys A
Once you have a trichomycete in front of you, how do you know what it is?
The best place to quickly identify your tricho is to navigate to the Lucid Keys
You will see a list (A) to choose from
If you have a fungal species, the choices highlighted in pink are the ones to examine. If you’re dealing with a protist, choose one of the other links.
Once you’ve clicked on your link, a screen appears (B) where you can input measurements, hosts, gut attachment, etc. and retreive a list of species that match your features.
This will all become more clear as you go through the handbook and familiarize yourself with trichos. We will revisit this sight in Part 5 – Morphometrics. B

8. Overview of Trichomycetes
Part 1
Overview of Trichomycetes
Insert pictures
Harpellales
Asellariales
Amoebidiales
Eccrinales

9. Overview*
Trichomycetes are obligate endosymbionts of a variety of arthropods.
Their hosts may be immature aquatic stages of insects, as well as adult terrestrial arthropods.
Trichos reside in the digestive tract of these organisms commensally – that is, they have a neutral relationship with their host. However, there have been several studies showing that they can shift between all aspects of symbiosis including mutualism and parasitism.
There are currently 385 species of trichos (including both fungal and non-fungal members).
Their systematics is in a state of flux, but molecular phylogeneticists are in the process of stabilizing this group of organisms in order to strengthen the base of the fungal tree of life.
*This section is decidedly brief. Refer to the monograph and further reading (Appendix III) for more information.

10. Overview - Symbiosis Shifting continuum
Symbiosis refers to two or more organisms living in close association with each other. It spans from parasitism through commensalism to mutualism.
Interestingly, trichomycetes exist along the entire spectrum of symbiosis.
Shifting continuum
Symbiosis
Mutualism
Commensalism
Parasitism
Host
Symbiont
Culisetae Studies on the Nutritional Relationship of Larval Aedes aegypti (
Smittium culisetae (Trichomycetes)
Author(s): Bruce W. Horn and Robert W. Lichtwardt
Source: Mycologia, Vol. 73, No. 4 (Jul. - Aug., 1981), pp
Morbosum
Sweeney, A. W. 1981a. An undescribed species of Smittium (Trichomycetes) pathogenic to mosquito larvae in Australia. Transactions of the British Mycological Society 77: 55-60
Host
Symbiont – +
Host
Symbiont ø +
Effects
Tricho example
Smittium morbosum§
Default for all other trichos
Smittium culisetae ¥
¥ Horn & Lichtwardt, 1981, Mycologia 73:
§Sweeney, 1981, Transactions of the British Mycol Soc, 77:

11. Overview – Taxonomy
The Class Trichomycetes were described by R.W. Lichtwardt in 1954.
All members were originally described as fungi (trichomycete translates to “hair fungus” because of the hair-like nature of the organisms in the gut)
Traditionally there were four orders described: Amoebidiales, Asellariales, Eccrinales and Harpellales.
In 2002 it was confirmed by Mendoza¥ that the Amoebidiales were non-fungal, and are now considered ichthyosporean protists.
In 2005, using molecular phylogenetics, Cafaro§ found that the Eccrinales were also protists.
The Harpellales and Asellariales are true fungi, yet their phylogenetic placement as early diverging members of the fungal tree of life is in flux.
Currently, scientists are resolving the issues of many clades and finding new relationships among taxa using new molecular technology.
Later in this guide you will learn how to preserve a fungal specimen so that its DNA can be extracted for analysis.
Get references for amoebidiales and eccrinales.
Mendoza, L., J.W. Taylor, and L. Ajello The Class Mesomycetozoea: a heterogeneous group of microorganisms at the animal-fungal boundary. Annual Review of Microbiology 56:
¥Mendoza, Taylor & Ajello, 2002, Ann Rev Microbiol, 56:
§Cafaro, 2005, Molecular Phyl & Evoln, 35: 21-34

12. Taxonomy Fungi Protists Order Host Asexual Spore Sexual Spores
Here is a table of some non-genetic features that separate the fungi from the protists
Order
Host
Asexual Spore
Sexual Spores
Fungi
Harpellales
Larval aquatic Insecta
Trichospore
Zygospore (biconical)
Asellariales
Isopoda or Collembola
Arthrospore or Sporangiospore
Zygospore (round)
Protists
Eccrinales
Diplopoda, crustacea, Insecta
Sporangiospore
N/A
Amoebidiales
Aquatic Crustacea or Insecta
Cystospore

13. Collecting Trichomycetes
Part 2
Collecting Trichomycetes
Pictures are from sampling at Dry Creek, February, From left to right: Prasanna Kandel, Emma Wilson, Nicole Reynolds, and Zach Hoefer.

14. Collecting Trichomycetes
It is a relatively simple process to collect trichomycete hosts from nature
Although there are many hosts of trichos, this section will focus on methods for collecting one of the true fungal members of the group: Harpellales*
We’ll provide locations that hosts are likely to be found, a list of supplies needed for aquatic sampling, and methods on how to sort the “good” from the “bad” aquatic insects.
pictures
*Whenever you collect Harpellales, non-fungal members in the genus Paramoebidium are often found in the same hindgut

15. Where are trichos found?
Where aren’t they found?!
Most immature aquatic insect hosts require flowing water, so streams are a great place to find hosts.
Aquatic hosts are found everywhere from pristine streams (A) to irrigation ditches (B)
Often, shed skins (exuvae) of hosts contain trichomycetes, so collecting in pools where they’ve washed up may also prove fruitful (C). B
Reynold’s Creek, Idaho
Sarah Oman in Parma, Idaho C
Emma Wilson and Yan Wang, Boise River, Idaho

16. Collecting Freshwater Aquatic Hosts
Prasanna Kandel showing off his hip waders!
For sampling aquatic hosts you’ll need the following items:
Boots (hip or chest waders work well)
Life vests for deep water
Nets (D-nets usually work best)
Buckets (optional)
Resealable plastic bags and/or tupperware containers
Transfer pipettes and forceps
100mm Petri dishes (for black fly larvae)
Plastic pans
Stream thermometer
Cooler with ice for transport to the lab

17. Collecting Freshwater Aquatic Hosts
When sampling from lotic habitats (flowing water), its best to stand upstream from your net and use one of your feet to overturn rocks, kick leaves, and disrupt the area to bring insects up from the bottom.
Then, swish your net in a figure eight to capture all of the debris that is floating downstream (including insects!)
Put a bit of water in your pan, then shake your net into it.
You can then sort the desired insects from the large pan into your small tupperware containers, or simply pour your pan into the plastic bags and sort in the lab.
Put your tupperware or plastic bags on ice and head back to the lab.
Yan Wang collecting hosts in the Boise River
Emma Wilson sorting insects with students at Reynolds Crk
Prasanna Kandel and Lance Steele sorting insects at Cottonwood Crk
Sarah Oman and Emma Wilson packing samples into the cooler in Parma, ID

18. Collecting Black flies (Simuliidae)
Black flies are lower dipterans that are a great host of trichos (see guide in Appendix II).
Depending on stream conditions, you may not capture all of the available black flies because they tend to cling to substrates such as dangling vegetation
Therefore, it helps to swish the net through or pull up manually any trailing vegetation.
They may also be attached to rocks and boulders, so bringing them out of the stream and hand picking is also useful.
Mass of black fly larvae on a twig from Dry Creek, Idaho

19. Sorting Aquatic Hosts
The Good
The Bad
So now you have your insects in the pan – which ones do you take back to the lab?
In Appendex II you can find keys to identifying hosts to the family level.
The “bad” stonefly pictured above is in the family Perlidae, which are predaceous.
Not only are trichos not found in predaceous insects, but the good hosts (like the Nemouridae above) could be eaten by the undesirable hosts.
Hmm... Which ones to take?
Dr. White ponders which insects to collect … and perhaps the meaning of life?

20. Sorting Aquatic Hosts
Sorting hosts may be done in the field or in the lab.
The keys in Appendix II will help to identify the correct (i.e. non-predaceous hosts)
Using your pipette, transfer insects from the large collecting pan to the smaller tupperware pans, partially filled with stream water.
If the host is too big for the pipette, use the feather-tipped forceps to move them.
Black flies have very high oxygen demands, so if they are to be placed in deep water, they don’t survive as well (unless the pan is aerated upon return to the lab).
Therefore, it helps to place black fly larvae in petri dishes with a very thin film of water over them (see bottom picture).
This way, they can still remain moist, and have enough gas exchange through the small amount of water.
From left to right: Mayfly, stonefly, black fly, midge
Pictures of various hosts: mayflies, stoneflies, black flies, midges, caddis flies, etc. – show ex. Of good hosts.
Emma Wilson sorting hosts with high school kids at
Reynold's Creek, Idaho

21. Maintenance, Dissection and Preservation
Part 3
Maintenance, Dissection and Preservation

22. Maintenance, Dissection and Preservation
A good deal of work in the morphology lab deals with maintaining, dissecting, and preserving hosts and trichos.
This section will guide you through this process in the following sections:
Documentation of samples
How to keep hosts alive in the lab
Dissection example and explanation
Preservation of host
Preservation of trichos (slides and CTAB)

23. Documentation of Samples
For each collection event, a unique collection code should be made as follows, separated by a hyphen:
ID-55
State, Province or Country code (usually 2-3 letters)
Serially increasing site reference number. May be accompanied by a letter to designate sub-sites in the location (e.g. “ID-55A”)
The sample code should be recorded in your field and lab books, or collection log, along with any relevant site information (stream temp, vegetation, substrate, GPS).

24. Maintaining Hosts in the Lab
The last thing you want, is for your sample to quickly die after returning to the lab.
The best way to prevent this is to dissect your hosts as quickly as possible.
Another approach, detailed here, is to create livable conditions for them in the lab itself.
It helps to keep black flies in a petri dish with a thin water overlay for short term storage, as previously described.
Take new pictures
An advanced setup with manifolds and airlines pumped through air diffusers to maintain hosts in the lab.

25. Maintaining Hosts in the Lab
For long term storage of hosts, a simple aeration system can be implemented.
You will need:
Air-diffusing stones
Air tubing
Air pump
Tupperware containers
Cut a small hole in the top of the tupperware and run a hose through, connected to an air pump.
Keep the sample in the fridge with the air pumping through the stone.
If the food looks depauperate, they can be fed a fish-food slurry (ground up tetra-fin in water).
Add pic from phone
Tupperware container
Air pump
Tubing with airstones attached

26. Just before dissection…
They’re often really small!
Keep your hosts on ice in either a tupperware container or petri dish.
This will keep them cool and calm.

27. Dissecting Some useful supplies
For the most complete dissecting experience, you will need the following:
Stereo-microscope (one with dark-field lighting is best)
Glass slides (3”x1”x1.0mm)
Coverslips (glass 18x18 mm is best)
Fine tipped jeweler’s forceps
Insect mounting needles held by pin vices
Droppers of
Distilled Water
Lactophenol Cotton Blue (LCB)
NanoPure water
Small vials for host specimens
Also, small pieces of paper for vials
CTAB [Cetrimonium bromide ((C16H33)N(CH3)3Br] – a DNA buffer
70% Ethanol (EtOH)
Delicate task wipers
A lab notebook
Pen and pencil
Some useful supplies h g c b a i e d f k j l

28. Dissecting
To begin with, take a clean glass slide and place two drops of water on it
The one on the left will be your “kill drop”
This is where you’ll be dissecting the gut out of the host
The one on the right will be your “dissection drop”
This drop may be preserved for future identification
Therefore, don’t put the drop too far to the right, otherwise it may not image very well
Place your host in the “kill drop” and begin dissecting.
“kill drop”
“dissection drop”

29. Dissecting – Dipterans
We’ll be using a black fly to demonstrate how to dissect a dipteran host.
Step 1 – Remove head and anus A A C B
Using one forcep, grab some exoskeleton on the section just below the head capsule – try to pin the larva down. With the other forcep, pinch the area to cleanly remove the head.
The anus can be seen by an “X” on the dorsal side of the larva. Its usually best to penetrate the insect with forceps and tear the anus open.
Look for the peritrophic matrix (clear tube coming from the body cavity – yellow star above, white arrow in facing picture). Often, the hindgut will protrude from the anal opening (black star above). B C

30. Dissecting – Dipterans: Midgut
The intact midgut
foregut 1.
Peritrophic matrix with food inside 2. D E F G
Lifting PM to remove food, using needle 3.
D. Once the gut is exposed, you can begin to remove the midgut, or peritrophic matrix (PM)
The peritrophic matrix is a clear “tube” that lines the inside of the midgut.
At the top, you can often see the foregut (image 1).
No gut fungi are found in the foreguts of dipterans, so it is best to remove the clear lining, which is often filled with food (image 2).
To remove the food, slide the needle underneath the PM, lift and lower it until it is as empty as you can get it (image 3). You may need to add more water to your slide to get all of the debris out.
Once the food is out, place the PM in a new drop of water (usually on a new slide) to image the gut fungi. Under the dissecting microscope, the trichos appear as small clear hairs (image 4).
Midgut fungi observed as tiny hyaline hairs 4.

31. Dissecting – Dipterans: Hindgut1. D E F G
The intact hindgut 2.
F. To isolate the hindgut, detach the gut just posterior to the malpighian tubules, as pictured above with the black arrow (F).
The intact hindgut (image 1) is a coiled gut that is slightly opaque.
- Place the hindgut in the dissection drop, and use the needles to tear the gut open and into smaller pieces to release all of the trichos - both fungal and protistan symbionts may be present (image 2).
The dissection drop is often full of food, debris, and “host bits”. The example specimen had a Paramoebidium (non-fungal associate) in the hindgut - pictured in the circle (image 3).
If a tricho is found, you may mount it on a separate slide by picking up the fungus with the needle (similar to how the PM was picked up on the previous page) and placing it in a new drop.
Using needles to tear open the hindgut and expose fungi 3.
Post-dissection. Can you see the tricho in the circle?

32. Dissecting – Non-Dipterans
We’ll be using a mayfly (Ephemeroptera, Baetidae) to demonstrate how to dissect a non-dipteran host. A A B C B D C
The intact hindgut D
Non- dipteran hosts (such as stoneflies and mayflies) do not require dissection of the midgut.
They are dissected in a similar way to dipterans as far as head removal (A), but the hindgut may be dissected out by grasping the abdominal segment just anterior to the rectum and pulling (B), exposing the hindgut and malpigian tubules (arrow, C). The gut may then be dissected in the same fashion as described in the previous section (D) using the needles.
One thing to be careful of is the extrusion of some species of trichos outside the anus (yellow star, C). When this happens, take extra care to not disrupt them as they often have produced sexual spores. You can place the entire anal segment on the slide, or try to remove them with your forceps and make a new slide.

33. Slide Preparation
ID-55
State, Province or Country code (usually 2-3 letters)
Serially increasing site reference number. May be accompanied by a letter to designate sub-sites in the location (e.g. “ID-55A”)
Now that you have your tricho on a slide, what do you do with it?
Carefully place the coverslip on the slide, avoiding air-bubbles and having the specimen squish out from underneath. If your slide is really wet, wait a few minutes for it to dry before proceeding.
Using a slide-specific marker, write the collection code followed by your initials and the number you’re on.
For instance, in the slide pictured, it was collection number 145 in Idaho (ID-145), and the dissector’s (Emma) first host (E1).
Also write:
Host ID (common name is fine)
Date
What is on the slide (midgut, hindgut, dissection drop)
The genus and species (if you know it)

34. Slide Preparation A
After writing the description on the left side of the coverslip, add a tiny drop of lactophenol cotton blue (LCB) to the right side of the coverslip.
As shown in the top panel of images (A), its best to completely empty the dropper of LCB to get a small droplet at the base.
This can then be applied to give just the right amount (B).
Next, apply a coat of clear nail polish (we use Revlon Clear, #771) over the text so that it doesn’t wash away while prepping the slide (C).
The LCB is then allowed to infiltrate the coverslip (D) for a few minutes to days.
The slide is kept flat (don’t ever tilt the slide until it is completely fixed!) on a letter tray while it is being prepared (E) . B C D E

35. Slide Preparation A B
The LCB is completely infiltrated when blue can be seen through the whole coverslip (A).
To wick off any excess LCB, a small piece of Kimwipe can be placed on the right side of the coverslip where the LCB drop was originally placed (B).
This can be left for a few hours to days.
Once the excess has been removed, clear nail polish is applied around three sides of the coverslip (C)
The right side is left bare (where the LCB was applied) (D) so that it may be rinsed in the next step. C
Right side is left bare D

36. Slide Preparation
Once the nail polish has dried, the excess LCB needs to be rinsed off with distilled water.
Make sure that the slide is tilted to the right so the LCB rinses away from the text (see image).
After the slide dries, apply nail polish to all 4 sides of the coverslip.
Nail polish should be applied until there is a smooth transition between the slide and coverslip
You should be able to rub your finger along the nail polish and not be able to feel the sharp margin of the coverslip.
Once it is completely sealed, the slide can be put away in the appropriate slide box, and saved for future inspection.
Store the slide with the coverslip facing UP so you are working with gravity!
Remember, the more thorough the slide preparation, the longer the slide will last!
Apply to all sides after rinsing

37. Moist Chamber A
Sometimes when you have a tricho, it helps to place the slide in a moist chamber to facilitate detachment of spores and the development of other features.
You will need the following items (A):
Filter paper – Petri dish
Distilled water – Bent glass rod
To make the moist chamber…
Place the filter paper in the petri dish (B)
Squirt a small amount of water into the dish (C)
Place glass rod on top of filter paper (D, E)
Put slide (with coverslip on and some extra water on the right side of coverslip) on top of the bent glass rod (F).
Cover petri dish, and label the isolate number, what is on the slide, date and time you made the moist chamber (G).
After a few hours (usually no more than one day) come back and image the slide to see if the spores have detached.
The high humidity in the chamber allows us to manipulate the tricho in ways that are otherwise impossible. B C D E F G

38. CTAB – Preservation of DNA
CTAB is a buffer used to preserve DNA prior to extraction for genetic analysis in the molecular lab (details not included in this handbook).
Clean the dissecting microscope with 70% ethanol, and wear gloves. Take CTAB out of refrigeration and allow to thaw (a few minutes until the white pellet is not visible).
Remove as much of the host tissue as possible, and move the fungus to a clean slide with a drop of NanoPure water (free of DNA) (A).
Clean your needles thoroughly first with 10% Bleach (B) by completely submerging the needle and part of the pin vice (C).
Repeat with 100% ethanol (D) and allow to dry before continuing.
Carefully open the top of the CTAB container with your pinky – don’t touch the lip of the jar (E).
Carefully pick up the specimen and place inside container, while holding the lid facing down (arrow, image F).
Write the slide code on the lid of the vial (G) and the contents on the side of the vial (H). C D E F G H

39. Host Preservation A
It is important to save the hosts after dissection for identification purposes
Fill a small vial with 100% ethanol (A)
Write the slide code on a small piece of paper (1-2 cm long) using pencil (B) and put in vial
If you use pen, the writing will bleed off in the ethanol
Pick up host with forceps and place in vial (C).
You now have a preserved host! (D)
The vials are then stored in boxes, and the information is entered into a spreadsheet for easy look-up. B C D

40. Part 4
Culturing

41. Culturing
Culturing trichomycetes has its challenges – especially given the obligate symbiotic nature of these organisms.
However, methods have been formulated for successful culturing of about 1/3 of the species of trichomycetes
This section will provide:
Media recipes (including modifications for slants, plates and liquid shake culture)
How to make media
How to pour plates and slants
How to transfer cultures between slants.
pictures

42. Recipe: 1/10 BHI Plates: Water: use 1 L per 1 L BHI
BHI: use 3.7 g per 1 L BHI
Agar: use 15 g per 1 L BHI
Slants:
Agar: use 19 g per 1 L BHI
Broth: NO AGAR
BHI is short for Brain Heart Infusion
Provides good media to obtain prolific sporulation
It’s the easiest recipe for tricho culturing and only has 3 ingredients: BHI, Agar and water.

43. Recipe: TGv – stock solutions
Biotin/Thiamin – 50 x solution: use 20 ml per 1 L TGV
to make 50x Biotin/Thiamin solution
0.25 mg Biotin/100 ml water
1.0 mg Thiamin/ 100 ml water
Mix these together to get 50x solution
use 20 ml of the 50x solution per 1 L TGV
Calcium Chloride – 50x solution: use 40 ml per 1 L TGV
to make 50x Calcium Chloride solution
0.175g CaCl2 /100 ml water
KEEP SEPARATE FROM SALT SOLUTION – WILL PRECIPITATE OUT IF MIXED
use 40 ml of the 50x solution per 1 L TGV
Salt Solution – 10x solution: use 100 ml per 1 L TGV
to make 10x Salt Solution add the following to 500 ml water:
0.7 g KH2PO4
0.875 g K2HPO4
0.65 g (NH4)2SO4
0.25 g MgCl2. 6H2O
use 100 ml per 1 L TGV
TGv is short for Tryptone Glucose and vitamins
This is slightly more complicated, and requires the formation of stock solutions to keep around for ease of culturing in the future.
This media produces excellent growth of the thallus and is ideal for building up the mass of a culture.
Maybe break this up and show pictures

44. Recipe: TGv – Tryptone Glucose vitamins
Plates:
Water: use 1 L per 1 L TGV
Agar: use 15 g per 1 L TGV
Glucose: use 5g per 1 L TGV
Tryptone: use 20 g per 1 L TGV
Slants:
Agar: use 19 g per 1 L TGV
Glucose: use 5 g per 1 L TGV
Broth: NO AGAR
Biotin/Thiamin – 50 x solution: use 20 ml per 1 L TGV
Calcium Chloride – 50x solution: use 40 ml per 1 L TGV
Salt Solution – 10x solution: use 100 ml per 1 L TGV
More pictures

45. Recipes - BHIGTv
Plates:
Water: use 1 L per 1 L BHIGTv
BHI: use 1.85 g per 1 L BHIGTv
Agar: use 15 g per 1 L BHIGTv
Glucose: use 2.5 g per 1 L BHIGTv
Tryptone: use 10 g per 1 L BHIGTv
Slants:
Agar: use 19 g per 1 L BHIGTv Glucose: use 2.5 g per 1 L BHIGTv
Broth: NO AGAR
This media is the best of both worlds - it is half BHI and half TGv.
BHIGTv* stands for Brain Heart Infusion Glucose Tryptone vitamins
The stock recipes are the same, but the amount used is half that of TGv.
Biotin/Thiamin – 50 x solution: use 10 ml per 1 L BHIGTv
Calcium Chloride – 50x solution: use 20 ml per 1 L BHIGTv
Salt Solution – 10x solution: use 50 ml per 1 L BHIGTv
More pictures
*(pr: “big TV”)

46. Making media – Plates
Before making ANY media, be sure to coordinate with the stock room staff to ensure autoclave availability and timing!
Weigh ingredients and pour into 2 L Erlenmeyer flask
Add 1.0 L dH2O
Place flask on stirring hot plate (make sure there’s a stir bar in the flask)
Turn to lowest setting for stirring and heating
Stir for a few moments to get the “chunks” out of the bottom
No need to completely melt the agar for making plates
Cover top with a bit of aluminum foil and put on autoclave tape
Bring to the autoclave room
Authorized staff will have to start the autoclave for you, but you may be responsible for the timing of taking it out! (~45 min to 1hr)
After autoclaving, bring back to lab, and place on hot plate
Stir (no heat!) to maintain homogeneity
You are now ready to pour your plates!
Autoclave tape before being autoclaved (white stripes)
Break it up and show pictures
Note: this picture has water in the flask for demonstration and is NOT what the media should look like!

47. Pouring Plates Pictures and videos of demonstration coming soon…
Sterilize hood by spraying with 70% ethanol
Turn on Bunsen burner with sparker.
Keep plates and media flask near flame (zone of inhibition)
Pour molten media slowly, just filling the bottom of the plate.
If the media sloshes out of the bottom, set aside and keep pouring - they can be eliminated after all plates are made.
Flame the lip of the flask every few pours to maintain sterile environment.
Pour all plates until the media is used up
Keep in clean hood overnight to ensure no contamination

48. Making media – Slants
Before making ANY media, be sure to coordinate with the stock room staff to ensure autoclave availability and timing!
Weigh ingredients and pour into 2 L Erlenmeyer flask
Add 1.0 L dH2O
Place flask on stirring hot plate (make sure there’s a stir bar in the flask)
Turn to lowest setting for stirring and heating
Bring to a boil to melt agar then stir until ready to dispense
This may take a while, so be patient!
Once its homogenous:
Use the media dispenser in the stock room to fill 6 ml into each tube
Its nice to have someone help you put the caps on after filling
The caps should be loose before autoclaving to ensure sterilization
Put autoclave tape on rack (A)
Bring to the autoclave
Authorized staff will have to start the autoclave for you, but you may be responsible for the timing of taking it out
After autoclaving, bring back to lab and place rack on its side on 4 or 5 tongue depressors taped together (B, C)
This is the perfect height for the media to slant and not spill out
Allow to completely cool (overnight is best)
Once cool, tighten caps and refrigerate
Before use, put ~ 1mL sterile dH2O in each slant (D)
Use a new sterile pipette for each tube
Flame each tube before and after filling
Leave in clean hood overnight to make sure there is no contamination
You are now ready to grow some fungi! A B
Autoclave tape after being autoclaved (black stripes) C D
Sterile di H2O
Break it up and show pictures

49. Before making ANY media, be sure to coordinate with the stock room staff to ensure autoclave availability and timing!
Weigh ingredients and pour into 2 L Erlenmeyer flask
Add 1.0 L dH2O
Place flask on stirring hot plate (make sure there’s a stir bar in the flask)
Turn to lowest setting for stirring and heating
Stir until the liquid is homogenous
Pour desired amount of media into each media flask
Place cheese-cloth-wrapped non-absorbent cotton plug into each flask
Cover tops (white arrow on picture) with aluminum foil and put on autoclave tape
Bring to the autoclave room
Authorized staff will have to start the autoclave for you, but you may be responsible for the timing of taking it out!
After autoclaving, bring back to lab, and place in incubator
You are now ready to grow some fungi!
Making media – Broth
Break it up and show pictures
Note: this picture has water in the flask for demonstration and is NOT what the media should look like!

50. Culturing – Transferring Specimens between tubes
Its important to label all of your tubes with the appropriate isolate name, date of transfer, and media used.
Place the label on the new tube, and double check that the correct isolate is being transferred throughout the process.

51. Culturing – Transferring Specimens between tubes
Just as when pouring plates, its important to sterilize the hood:
Spray with 70% ethanol and wipe down
Also apply to your hands
Turn on Bunsen burner with sparker.
Let ethanol dry first
Keep tubes close to flame (zone of inhibition)
Start by flaming the loop holder (usually a small beaker, yellow circle, image A)
Flame the loops by dipping in 100% ethanol, then placing in flame.
Observe them turning bright orange (B).
Make sure the entire length of the loop gets flamed – anything going into the slant needs to be sterile.
Place loops on beaker and allow to cool before entering the slant. A B
Insert picture or video along with explanation.

52. Culturing – Transferring Specimens between tubes
Pick up the original slant with your non-dominant hand, and open the cap with your pinky on your dominant hand
Don’t touch the lid or the inside of the cap!
Once open, flame the lip by rotating for several seconds in the flame
Insert loop into slant and pick up a piece of fungus.
Hold fungus on loop (near zone of inhibition) while you re-flame the lip of the tube.
Place cap back on sterile tube
Be careful not to burn your hand or drop the fungus on the loop!
Retrieve new slant, open, flame and hold the same way as the original slant
Double check label for correct isolate number
Place loop in water at bottom of slant and break up fungus with loop-head.
Re-flame and close as before and place slant in tube rack
Ethanol and flame loop and place on loop-holder.
Keep tube caps slightly open (~1 twist) to allow for gas exchange.
While the fungus is growing, monitor daily by gently shaking and swirling water onto fungus
Tighten cap beforehand, and re-open after maintanence
More pictures and videos
of demonstration
coming soon…

53. Part 5
Morphometrics
Alison Chamberlin measuring Harpella thalli and spores.
Detached and measured spore of Stachylina

54. Morphometrics
One of the most important steps in dealing with trichos is accurately determining the identity of the species your working with.
This is valuable for documenting new locales of species, confirming species identification prior to molecular examination and tree building, and describing new species based on the features you’re observing.
This section of the guide will describe:
Tips for imaging
What and where to measure on your specimen
Example spreadsheet and what to include

55. Morphometrics - ImagingB
Morphometrics - Imaging
The Nikon 80i Eclipse A C
To image a slide, use the lab’s Nikon 80i Eclipse compound microscope (A).
The eyepieces are adjustable up and down, in and out (B)
The focus is on the left side (C)
The stage is moved with the knobs on the right (D).
Slides are placed on the stage as shown in image E.
BE SURE TO GET TRAINING CLEARANCE FROM A SENIOR LAB MEMBER PRIOR TO OPERATING THE MICROSCOPE. D E

56. Morphometrics - Imaging
Use the image software Spot Advanced (ver. 4.6).
Begin by opening the program while your slide is on the stage.
Click “Live” (arrow, A) to see the image on the microscope.
Adjust the focus manually
Capture the image by clicking the small camera with the red arrow above it (arrow, B).
Save the image by clicking File>Save As, and saving in the appropriate folder.
When saving, be sure to include the objective (i.e. 10, 40, or 60x) and slide code.
Every image is saved with a serially increasing unique number
All details are saved in the photo-log, located near the microscope. B

57. Morphometrics - Measuring
To add measurements to an image, open Spot Advanced
Click on Edit > Add/Edit Measurements
Select the appropriate magnification (in the example, Nikon 60x)
In the new measurement window, select the type of line you want to use (in this example, “straight line”) and click Create.
Position the “x” over the end of the spore (or whatever the object) and drag the line over the length of the spore.
A curved line can also be used to measure non-straight features
Such as the Harpella spore pictured on the bottom.

58. Morphometrics - Measuring
Where to measure:
By default, measure the widest area on the tricho feature you’re interested in
For example, the Stachylina to the right has measurements from the base to the tip of the spore (length), the widest part of the middle (width), and the length and width of the collar.
Morphometrics are quite variable between species
A list of features to measure and/or note are provided next.
Spore size
Collar size

59. Morphometrics - Measuring
Collection Slide
Code Status
Sub-site
Date
Pic#
On Slide
Host
Host stage*
Host Voucher? Selected for ID
Comments on final check
Fungus Genus Fungus Species
Fungus Development
T-Spore L1
T-Spore W1
T-Spore L2
T-spore W2
Z-Spore L
Z-Spore W
Z-spore angle
Z-ophore L
Z-ophore W
             
Here is a laundry list of features to include in your morphometrics table.
Some of these are things to actually measure (i.e. length and width) while others are comments or types (i.e. holdfast types)
All of these features will help you enter appropriate date into Lucid Keys (see beginning of handbook) and retrieve a species ID for your tricho.
Thallus L
Thallus W
Holdfast
Holdfast Length
Holdfast Width
HF Grade Change (L)
Basal cell length
Basal cell width
T/Z-spore shape
T-spore detached
T/Z-sp collar length
Density w/in gut
# t-spores/thallus
Generative Cell Length
Gen Cell Width
# appendages
cyst diameter
Oddities Comments              

60. Morphometrics - Measuring
Above is an image of a working morphometrics table.
All of the features from the previous slide are in the top row.
Numerical data such as t-spore length (T-Spore L1, see inset) are entered in a single cell, separated by commas.
This way, you can easily take the average, min, max and median of the data in Excel.
Qualitative data, such as fungus development (see inset) are written out and are useful for seeing non-quantitative trends.

61. Conclusions The tricho take-away It’s really not so bad…
Why is this so painful!

62. Consider the following:
It takes time to get comfortable identifying hosts in the field – don’t worry if you can’t ID them immediately. If you accidently dissect a predaceous stonefly it might actually help you see the difference between the gut contents.
Everyone has their own dissection techniques. The style described in this handbook are the author’s own bias and may not be ideal for all lab members.
Take your time, explore and experiment.
Don’t underestimate the importance of fixing your slide well. If done so carefully and correctly, they can last many years.
Morphometrics can take a very long time, but like most things in life gets easier the more you do it. Its very helpful to have an organized spreadsheet to work with – ask a senior lab member for a template to get you started!
The culture collection at Boise State University is the largest reserve of trichomycetes in the world. Continuous maintenance is crucial for the legacy of studying this group of organisms.

63. Thank you!
Funded by NSF Award
to Merlin M. White
DEB
This handbook was made for a graduate student project for the course Mycology (BOT330G) at Boise State University.
Special thanks to Eric Johnson and Justin Gause (undergraduate research assistants) for help with imaging and demonstrating techniques for this handbook.
Updates will surely follow, but in the mean time I sincerely hope that this will benefit trichomycete students for years to come.
GO TEAM TRICHO!!

64. I – List of genera II – Host keys III – Further reading
Appendices
I – List of genera
II – Host keys
III – Further reading

65. Appendix I – List of genera
This appendix can act as a quick reference to get the genus of your tricho
Remember…
Harpellales are in immature aquatic insects (for the most part)
The family Harpellaceae is in the midgut, Legeriomycetaceae is in the hindgut
Asellariales are in the hindgut cuticle of Isopoda or Insecta (Collembola)
Amoebidiales are on the external cuticle (exoskeleton) or hindgut cuticle of aquatic Crustacea or Insecta
The genus Amoebidium is in many arthropods, including crustaceans, Paramoebidium is in aquatic insects
Eccrinales is hindgut or foregut of Diplopoda, Crustacea, or Insecta

66. Appendix I – List of Genera: Fungal trichos
Order Harpellales
Family Harpellaceae
Harpella, Carouxella, Harpellomyces, Stachylina, Stachylinoides
Family Legeriomycetaceae
Allantomyces, Austrosmittium, Baetimyces, Barbatospora, Bojamyces, Capniomyces, Caudomyces, Coleopteromyces, Dacryodiomyces, Ejectosporus, Ephemerellomyces, Furculomyces, Gauthieromyces, Genistelloides, Genistellospora, Glotzia, Graminella, Graminelloides, Klastostachys, Lancisporomyces, Legerioides, Legeriomyces, Legeriosimilis, Orphella, Pennella, Plecopteromyces, Pseudoharpella, Pteromaktron, Simuliomyces, Sinotrichium, Smittium, Spartiella, Stipella, Tectimyces, Trichozygospora, Trifoliellum, Zygopolaris 
Order Asellariales
Family Asellariaceae
Asellaria, Baltomyces, Orchesellaria     

67. Appendix I – List of Genera: Non-fungal trichos
Order Eccrinales
Family Eccrinaceae
Alacrinella, Arundinula, Astreptonema, Eccrinidus, Eccrinoides, Enterobryus, Enteromyces, Enteropogon, Leidyomyces, Paramacrinella, Passalomyces, Ramacrinella, Taeniella, Taeniellopsis
Family Palavasciaceae
Palavascia
Family Parataeniellaceae
Lajasiella, Parataeniella
Order Amoebidiales
Family Amoebidiaceae
Amoebidium, Paramoebidium

68. Appendix II – Host Keys
This appendix should be used to determine your hosts identity to the family level
Don’t forget – always voucher your host so you can go back and double check your identification!
Show picture of pickled host

69. Here’s a great key for identifying some common aquatic hosts:
Appendix II – Host Keys
Here’s a great key for identifying some common aquatic hosts:

70. Appendix III – Further Reading
For more information, explore Dr. Lichtwardt’s website
See beginning of handbook
A list of the world literature can be found here