Trichomycete Morphology Handbook

Report
The Gut Guide: A handbook
for the trichomycete
morphology lab
Emma R. Wilson
Boise State University
Research lab of Merlin White, PhD
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
2
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.
3
Web Resources
• In addition to this guide, the following links are vital for anyone
interested in trichos:
– http://www.nhm.ku.edu/fungi/
– http://www.nhm.ku.edu/fungi/Lucid%20Keys.html
– http://www.nhm.ku.edu/~fungi/Monograph/Text/Mono.htm
• 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.
4
Web Resources
• On the home screen of http://www.nhm.ku.edu/fungi/ , 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
5
Monograph
• 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.
Contents are a great
place to start navigating
Glossary contains many
tricho-specific definitions
6
Lucid Keys
•
•
•
•
•
•
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.
A
B
7
Part 1
Overview of Trichomycetes
Harpellales
Asellariales
Amoebidiales
Eccrinales
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.
9
Overview - Symbiosis
•
•
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
Effects
Tricho
example
Parasitism
Commensalism
Mutualism
Host
–
Host
ø
Host
+
Symbiont
+
Symbiont
ø
Symbiont
+
Smittium
morbosum§
Default for all
other trichos
Smittium
culisetae ¥
10
¥
Horn & Lichtwardt, 1981, Mycologia 73: 724-740
§Sweeney, 1981, Transactions of the British Mycol Soc, 77: 55-60
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 nonfungal, 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.
¥Mendoza,
§Cafaro,
Taylor & Ajello, 2002, Ann Rev Microbiol, 56:315-344
2005, Molecular Phyl & Evoln, 35: 21-34
11
Taxonomy
Protists
Fungi
Order
Here is a table of some non-genetic features
that separate the fungi from the protists
Host
Asexual
Spore
Sexual
Spores
Harpellales
Larval aquatic
Insecta
Trichospore
Asellariales
Isopoda or
Collembola
Arthrospore or Zygospore
Sporangiospore (round)
Eccrinales
Diplopoda,
Sporangiospore N/A
crustacea, Insecta
Amoebidiales Aquatic
Crustacea or
Insecta
Cystospore
Zygospore
(biconical)
N/A
12
Part 2
Collecting Trichomycetes
Pictures are from sampling at Dry Creek,
February, 2011. From left to right: Prasanna
Kandel, Emma Wilson, Nicole Reynolds, and
Zach Hoefer.
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.
14
*Whenever you collect Harpellales, non-fungal members in the genus Paramoebidium are often found in the same hindgut
Where are trichos found?
• Where aren’t they found?!
A
B
Reynold’s
Creek, Idaho
C
Sarah Oman in
Parma, Idaho
• 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).
Emma Wilson and Yan Wang,
Boise River, Idaho
15
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
http://www.bioquip.com/prod_i
mages/7512D-001-TwoPiece-HD-Aquatic-Net.jpg
http://www.escalemodels
.com/forums/uploads/mo
nthly_08_2011/post2198094235900%201312467
236.jpg
http://www.babiestravellite.c
om/mm5/graphics/subcat_i
mages_sq/Ziploc-Bags.jpg
http://img.alibaba.com/img/pb/477/433/36
2/362433477_209.jpg
http://www.tritechresearch.com/items/T3361-L.JPG
http://image.become.com
/imageserver/s1/901007
951-75-75-5-32/streamthermometer.jpg
–
–
–
–
–
http://c1903.r3.cf3.rackcdn.com/YI132O770_1.jpg
16
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
17
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
18
Sorting Aquatic Hosts
• 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.
The Bad
The Good
http://academics.smcvt.edu/Vermont_rivers/River%20sites/Cold%20Brook.htm
Hmm...
Which ones
to take?
Dr. White ponders which insects to collect … and
perhaps the meaning of life?
19
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.
http://www.lifeinfreshwater.org.uk/Sp
ecies%20Pages/Midge_Chironomus.j
pg.html
From left to right: Mayfly, stonefly, black fly, midge
Emma Wilson sorting hosts with high school kids at
Reynold's Creek, Idaho
20
Part 3
Maintenance, Dissection and
Preservation
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)
22
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).
23
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.
An advanced setup with manifolds and
airlines pumped through air diffusers to
maintain hosts in the lab.
24
Maintaining Hosts
in the Lab
•
•
For long term storage of hosts, a
simple aeration system can be
implemented.
You will need:
–
–
–
–
•
Air pump
Tubing with
airstones attached
Tupperware
container
•
•
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).
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.
26
Dissecting
•
For the most complete dissecting
experience, you will need the following:
a.
Stereo-microscope (one with darkfield 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
b.
c.
d.
e.
f.
a.
b.
c.
g.
Small vials for host specimens
a.
h.
i.
j.
k.
l.
Distilled Water
Lactophenol Cotton Blue (LCB)
NanoPure water
Some useful supplies
i
j
a
h
f
k
b
g
c
l
d
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
e
27
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”
28
Dissecting –
Dipterans
We’ll be using a black fly to
demonstrate how to dissect a
dipteran host.
Step 1 – Remove head and anus
A
B
A
C
A.
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.
B.
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.
C.
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
29
Dissecting –
Dipterans: Midgut
foregut
1.
D
The intact midgut
E
2.
Peritrophic
matrix with
food
inside
F
G
Lifting PM to remove
3.
food, using needle
D. Once the gut is exposed, you can begin to remove the midgut, or peritrophic matrix (PM)
E.
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).
4.
Midgut fungi
observed as tiny
hyaline hairs
30
Dissecting –
Dipterans: Hindgut
1.
D
E
F
The intact hindgut
G
2.
F. To isolate the hindgut, detach the gut just posterior to the
malpighian tubules, as pictured above with the black arrow (F).
G. 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?
31
Dissecting –
Non-Dipterans
A
We’ll be using a mayfly (Ephemeroptera,
Baetidae) to demonstrate how to
dissect a non-dipteran host.
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.
32
ID-55
Slide Preparation
State, Province or Country
code (usually 2-3 letters)
•
•
•
•
•
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:
–
–
–
–
Serially increasing site reference
number. May be accompanied by a
letter to designate sub-sites in the
location (e.g. “ID-55A”)
Host ID (common name is fine)
Date
What is on the slide (midgut, hindgut,
dissection drop)
The genus and species (if you know it)
33
Slide Preparation
•
•
•
•
•
•
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) .
A
B
D
C
E
34
Slide Preparation
•
•
•
•
•
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.
A
B
C
D
Right
side is
left bare
35
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
–
•
Once it is completely sealed, the slide can be
put away in the appropriate slide box, and
saved for future inspection.
–
•
You should be able to rub your finger along the nail polish
and not be able to feel the sharp margin of the coverslip.
Apply to all
sides after
rinsing
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!
36
A
Moist Chamber
•
•
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
Distilled water
– Petri dish
– 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
37
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).
A
C
B
D
E
F
G
H
38
Host
Preservation
•
•
•
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
–
•
•
•
A
C
B
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.
D
39
Part 4
Culturing
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.
41
Recipe: 1/10 BHI
•
•
•
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.
RECIPE
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:
Water: use 1 L per 1 L BHI
BHI: use 3.7 g per 1 L BHI
Agar: use 19 g per 1 L BHI
Broth: NO AGAR
Water: use 1 L per 1 L BHI
BHI: use 3.7 g per 1 L BHI
42
Recipe: TGv –
stock solutions
•
•
•
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.
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
43
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:
Water: use 1 L per 1 L TGV
Agar: use 19 g per 1 L TGV
Glucose: use 5 g per 1 L TGV
Tryptone: use 20 g per 1 L TGV
•
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
Broth: NO AGAR
Water: use 1 L per 1 L TGV
Glucose: use 5 g per 1 L TGV
Tryptone: use 20 g per 1 L TGV
44
Recipes - BHIGTv
•
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
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:
Water: use 1 L per 1 L BHIGTv
BHI: use 1.85 g per 1 L BHIGTv
Agar: use 19 g per 1 L BHIGTv
Glucose: use 2.5 g per 1 L BHIGTv
Tryptone: use 10 g per 1 L BHIGTv
Broth: NO AGAR
Water: use 1 L per 1 L BHIGTv
BHI: use 1.85 g per 1 L BHIGTv
Glucose: use 2.5 g per 1 L BHIGTv
Tryptone: use 10 g per 1 L BHIGTv
45
*(pr: “big TV”)
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)
–
•
•
No need to completely melt the agar for
making plates
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
–
•
Turn to lowest setting for stirring and heating
Cover top with a bit of aluminum foil and put
on autoclave tape
Bring to the autoclave room
–
•
Autoclave tape before
being autoclaved
(white stripes)
Stir for a few moments to get the “chunks”
out of the bottom
–
•
Making media –
Plates
Note: this picture has water in the flask for
demonstration and is NOT what the media
should look like!
Stir (no heat!) to maintain homogeneity
You are now ready to pour your plates!
46
Pouring Plates
•
•
•
•
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.
–
•
•
•
Pictures and
videos
of demonstration
coming soon…
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
47
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)
–
•
–
–
This may take a while, so be patient!
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
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)
–
C
D
Allow to completely cool (overnight is best)
•
Before use, put ~ 1mL sterile dH2O in each slant (D)
–
–
–
–
Sterile
di H2O
This is the perfect height for the media to slant and not
spill out
•
•
Autoclave tape after
being autoclaved
(black stripes)
Put autoclave tape on rack (A)
Bring to the autoclave
–
•
Turn to lowest setting for stirring and heating
Once its homogenous:
–
•
•
B
Bring to a boil to melt agar then stir until ready to
dispense
–
•
A
Making media –
Slants
Once cool, tighten caps and refrigerate
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!
48
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)
Making media –
Broth
– 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!
•
•
Note: this picture has water in
the flask for demonstration
and is NOT what the media
should look like!
After autoclaving, bring back to lab, and place
in incubator
You are now ready to grow some fungi!
49
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.
50
Culturing – Transferring Specimens between tubes
•
Just as when pouring plates, its
important to sterilize the hood:
–
Spray with 70% ethanol and wipe down
•
–
•
•
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.
–
–
•
Also apply to your hands
Turn on Bunsen burner with sparker.
•
–
A
B
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.
51
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
–
•
•
•
•
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 reflame the lip of the tube.
Place cap back on sterile tube
–
•
•
•
•
•
More pictures and videos
of demonstration
coming soon…
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
–
•
Don’t touch the lid or the inside of the cap!
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
52
Part 5
Morphometrics
Alison Chamberlin measuring Harpella thalli and spores.
Detached and measured spore of Stachylina
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
54
B
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
55
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.
A
B
56
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.
57
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.
Spore size
Collar size
• Morphometrics are quite
variable between species
• A list of features to measure
and/or note are provided next.
58
Morphometrics Measuring
•
•
•
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.
•
•
•
•
•
•
•
•
•
•
•
•
•
•
•
•
•
•
•
•
•
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
•
•
•
•
•
•
•
•
•
•
•
•
•
•
•
•
•
•
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
59
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.
60
Conclusions
The tricho take-away
Why is
this so
painful!
It’s really
not so
bad…
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.
62
Funded by NSF Award
Thank you!
to Merlin M. White
DEB-0918182
•
•
•
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!!
63
Appendices
I – List of genera
II – Host keys
III – Further reading
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
65
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
66
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
67
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!
68
Appendix II – Host Keys
Here’s a great key for identifying some common aquatic hosts:
http://www.dep.wv.gov/wwe/getinvolved/sos/documents/benthic/dichotomouskey.pdf
69
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
70

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