Everything
Microbiology Lab!
A Guide for Teaching Assistants
Caroline’s Instructions, Tips and Tricks on Almost Everything Related to
Working in UHCL’s Microbiology Lab with Dr. LaMontagne
1
Microbiology Lab TA Responsibilities
1. Prepare materials for lab each week, including...
・ Media (Agar plates, broth, culture tubes, etc.)
・ Reagents (Aliquoted into smaller 5 ml tubes, preferably eight tubes per lab section)
・ Fresh bacterial cultures
・ Additional materials (loops, sterile 1.5 ml tubes, pipettes, etc..)
2. Review PPTs for the upcoming lab exercise(s) (which I will provide).
3. Test out lab exercise(s) beforehand to make sure everything is working properly.
4. Assist in overseeing lab exercises and supervising undergraduate students.
5. Enter in weekly notebook grades to excel spreadsheet (which I will provide).
6. Help with scheduling student make-up labs.
7. Help keep track of materials low in stock (will share inventory spreadsheet).
8. Let Dr. LaMontagne or Caroline take a break every now and then :)
2
Spring 2023 Tentative Schedule
Overview of
Weekly Lab
Exercises
Subject
to
Change
Techniques in bold are
ones that students
MUST do!!!
Week
Date
Technique(s)
1
18Jan23
Course Objectives, Biosafety, Lab Notebooks
2
25Jan23
Pipetteman, Parfocal, Pouring Plates
3
01Feb23
Streak-plates, Simple-Stain, JOVE: Aseptic Techniques
4
08Feb23
MALDI-TOF: Ethanol Preps, Crystal Violet Serial Dilutions
5
15Feb23
MALDI-TOF: Formic Acid Extractions, Gram-Stain, JOVE: Centrifuge
6
22Feb23
Track Dilution (Practice), SIM
7
01Mar23
MacConkey, Mannitol Salts Agar, Citrate, Phenol Red
8
08Mar23
Catalase, Casein Hydrolysis, Starch Hydrolysis, Track Dilution (Practical)
9
13-19Mar23
Spring Break (NO CLASS)
10
22Mar23
Literature Search, Group Project Outline
11
29Mar23
EtNa DNA Extraction
12
05Apr23
16S PCR
13
12Apr23
Unknown ID, Kirby-Bauer
14
19Apr23
Continue Group Projects & Unknown ID
15
26Apr23
Continue Group Projects & Unknown ID
16
03May23
Finish Group Projects
3
Preparing
Materials for Lab
The following section includes…
・Protocols for making agar plates, culture tubes, and broth stock bottles
・Some tips & tricks for preparing media and pouring plates
・General instructions on preparing reagents for lab exercises
・Bacterial strains used in lab and preparing/maintaining cultures
・Biohazardous waste disposal
4
Preparing Materials for Lab
Preparing Materials for Lab
Essential Materials We Should Always Have Readily Available:
・ TSA plates (Preferably at least 15-20 plates at any given time)
・ Sterile 1x PBS
・ Autoclaved 1.5 ml microcentrifuge tubes
・ 70% Ethanol
・ Pipette tips (P1000, P200, P10)
・ Gloves (S, M, L, XL)
General Lab Materials We Need to Keep in Stock:
Regarding Amount of Plates
・500 mL should make around 20-25 plates.
・Assuming no one drops the course…
There’s 64 lab students/semester.
・3 flasks of 500-600 mL will make enough for each lab!
・We only have two hot/stir plates, so what I do:
・ Sterile loops
・ Conical tubes (50 mL, 15 mL, and 5 mL)
・Allow the 3rd flask with water to heat on a hot plate
We have tons (without stirring though).
Amount of Plates/Tubes per Exercise:
・Use autoclave glove to shake the hell out of it every now
and then while waiting for an available hot/stir plate.
・ For Petri dish exercises, we need one plate per student
・Letting powder settle will cause it to burn though!
・ For culture tube exercises, we need two tubes per student
- So, we’ll need to make around 140 tubes per media…
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Preparing Materials for Lab
General Media Composition
I. Water
・To dissolve materials to be transported across the cytoplasmic membrane
II. Carbon Source
・Required for the construction of all organic molecules
・Usually glucose but can be other carbohydrates depending on purpose, such as:
Important to know in general,
but especially since we tend to
make quite a few mediums
from scratch!
- Lactose, Sucrose, Mannitol, etc..
III. Nitrogen Source
・Needed for the synthesis of amino acids, DNA, RNA, and ATP, among other molecules
・Depending on the organism, nitrogen sources can include…
- Nitrogen, Nitrates, Ammonia, or organic nitrogen compounds
IV. Buffer System
・Proper pH VERY important for microbial growth
・pH buffer acts as either a weak acid or a weak base to ensure that the media will be somewhat
resistant to change in pH
V. Mineral Source
・Micronutrients, including minerals and vitamins, needed for bacterial metabolic pathways
6
Preparing Materials for Lab
Preparing Agar Plates
General approach to preparing nutrient agar plates.
Please read the instructions specific to the media you are making prior to beginning, as some media have
specific preparation requirements.
1. Add 500 ml of ddH2O to a 1000 ml flask and begin heating on hot plate.
・ Media must only fill ½ volume of flask
・ Cover with foil while heating to avoid water loss
2. Weigh out amount of dehydrated powder needed for a final volume of 500 ml
・ Bottle will tell you how many grams per liter, so divide that in half
3. Slowly add powder to flask while stirring on hot plate, and cover with foil.
4. Allow media to heat to a boil for approximately one minute to fully dissolve media.
5. Sterilize by autoclaving at 121∘C, 15 psi for 15 minutes.
6. Remove from autoclave and allow media to cool to 50∘C prior to pouring into Petri dishes.
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Preparing Materials for Lab
Preparing Culture Tubes
General approach to preparing agar or broth culture tubes.
1. Add 500 ml of ddH2O to a 1000 ml flask and begin heating on hot plate.
・Cover with foil while heating to avoid water loss
2. Weigh out amount of dehydrated powder needed for a final volume of 500 ml
3. Slowly add powder to flask while stirring on hot plate, and cover with foil.
4. Allow media to heat to a boil for approximately one minute to fully dissolve media.
5. Let media cool to approximately 55∘C and transfer 10 ml to each 15 ml glass culture tube.
・ Place plastic cap loosely on culture tubes
6. Sterilize by autoclaving at 121∘C, 15 psi for 15 minutes.
7. Remove from autoclave and allow media to cool slightly before tightening culture tube cap.
8. Allow media to cool at room temperature on the lab bench.
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Preparing Materials for Lab
Preparing Broth Stock Bottles
General approach to preparing stock bottles of broth.
1. Add 500 ml of ddH2O to a 1000 ml flask and begin heating on hot plate.
・Cover with foil while heating to avoid water loss
・The bottles we use to autoclave/store broths are not meant to be heated on a hot plate
2. Weigh out amount of dehydrated powder needed for a final volume of 500 ml.
3. Slowly add powder to flask while stirring on hot plate, and cover with foil.
4. Allow media to heat to a boil for approximately one minute to fully dissolve media.
5. Transfer the 500 ml of broth to a 500 ml screw-top bottle.
・ Place screw-top cap loosely on bottle prior to autoclaving
・ Bottles are designed to not boil over in autoclave, so filling full volume is fine
6. Sterilize by autoclaving at 121∘C, 15 psi for 15 minutes.
7. Remove from autoclave and allow media to cool slightly before tightening screw-top cap.
8. Allow media to cool at room temperature on the lab bench.
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Preparing Materials for Lab
Media Preparation Tips
1. Keep bottles of dehydrated media in closed sliding glass shelf
・Dehydrated media can be negatively effected by environmental factors,
regardless of the bottles being closed.
・Humidity is the factor that affects our dehydrated media the most.
・Aside from living in a humid place in general… We have our dehydrated
powder in B3213, where the autoclaves and warm water bath are kept.
・BOTH of which produce steam/water vapor!
・Water vapor will hydrate the powdered media, causing it to clump.
・To help avoid this, please store bottles of dehydrated media in the sliding
glass shelf (B3213) and close shut :)
10
Preparing Materials for Lab
Media Preparation Tips
2. Make 600 ml in a 1000 ml flask at a time
・ We normally make 500 ml at a time but 600 ml will not boil over in the autoclave.
・ So, I make 600 ml of TSA and other media for lab exercises at a time!
- Keeping in mind that media bottles include the g/L needed…
Just divided that by 10 to see how much you need for the extra 100 ml
- Ex. TSA calls for 40.0 grams per liter, so just move that decimal to the left
and you’ll see it’s 4 grams per 100 ml!
・ 100 ml can make around five extra plates, depending on thickness.
- Sometimes just five extra plates can be a life saver… (Many times, really...)
・ ‘Shortcut’ for us, not for undergrads. They must learn to only fill ½ volume of flask.
・ I’ve found that three flasks of 600 ml is perfect to make media for all four lab sections
in one go! Just be mindful to not let it boil over on the hot plate!
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Preparing Materials for Lab
Media Preparation Tips
3. Heat water prior to adding dehydrated powder
・ I always start heating the flasks of water prior to measuring out the powder.
・ Even getting it just a little warm is better than cold water.
– It clumps otherwise!
4. Keep flask covered with foil and crank that heat up to 300-320∘C
・ Unless medium requires specific temperature!! (ex. TCBS)
・ Water loss from evaporation will change the total volume of your media,
which can effect things like the concentration of agar, salt, etc..
・ Heating at 300-320∘C will not cause media to burn (unless agar clumps at bottom).
・ I see most TAs preparing media at ~200∘C, which is a waste of time.
Be mindful of the temperature media components degrade at,
but I’ve never had any issues in five years bumping up the temp this high for our lab media.
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Preparing Materials for Lab
Media Preparation Tips
5. If pressed for time, add powder to flask and shake before stirring
・Just add the powder to the flask with water, cover with foil, and shake the hell out of it by
hand a few times.
・Saves a TON of time and helps prevent agar clumping at the bottom/burning.
・Skim Milk agar MUST be shaken by hand before heating!
- Takes a long time to fully dissolve but will burn very easily, very quickly.
- Add a small amount of powder to flask, shake until no clumps, and
repeat with remaining powder. Then stir on hot plate.
Just a shortcut for us, really.
Students must learn proper
method of slowly adding
powder to water while
mixing.
・Always add powder to water! Not water to powder!! It will clump and burn!
・Always add powder to flask slowly, preferably a little at a time. Adding too fast can cause
powder to aerosolize.
・At some point, I’ll likely ask you to make a 5 mM CTAB solution and CAS-R2A agar... DO NOT SHAKE!
- CTAB is a detergent (soap) and will bubble/foam like CRAZY.
13
Preparing Materials for Lab
Media Preparation Tips
6. When making solid agar from a broth, start by mixing the agar first.
・ The agar we buy as of late comes in flecks instead of powder
・ Takes longer to dissolve so I like to dissolve it prior to mixing anything else!
・ Solid media can be made with agar concentrations of 1.0 – 2.0%
・ We use 7.5 g of agar per 500 ml of water, which makes 1.5% agar plates
・ “Soft” agar has concentrations of ~0.05 – 0.5% (Mostly for motility studies, ex. SIM)
・ If agar concentration is too high, motility is restricted and bacteria may have
to rely on nutrients diffusing to them (Resulting in less growth)
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Preparing Materials for Lab
Media Preparation Tips
7. When Making Broths:
・ Heat and dissolve in a flask as you would with agar.
・ The bottles we use for broths are not meant to be heated on a hot plate.
・ After fully dissolving and mixing, transfer broth to screw-top bottle before autoclaving.
・ Leave cap loosened while autoclaving (but tighten afterward!)
8. When Making Media for Culture Tubes:
・ Let the media cool a bit before adding it to the tube/covering the tube with a cap.
・ Letting it sit in a 50-55∘C water bath will help.
・ Excess condensation from heat can cause media like SIM or Citrate to not solidify properly.
・ Add approximately 10 ml of broth per tube prior to autoclaving.
・ Leave cap loosened while autoclaving (but tighten/push down afterward!)
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Preparing Materials for Lab
Media Preparation Tips
9. Not all media reagents can/should be autoclaved!
・ If you’re making a media from scratch using a recipe, make sure all of the
components can be autoclaved.
・ Solutions that contain heat-labile components must be filter-sterilized using a
0.22 µm filter.
・ Some compounds will degrade into other compounds when autoclaved, so they
must be filter-sterilized and added after other media has autoclaved and cooled.
・ Example. CaCl2 cannot be autoclaved in the presence of sulphates (such as MgSO4)
or it will precipitate as calcium sulphate.
・ It is best to autoclave glucose solutions alone if not filter-sterilizing.
16
Preparing Materials for Lab
Media Preparation Tips
10. Autoclaving medium components separately:
・Some components of media may be better to autoclave independently as they
interact with one another while autoclaving.
・A sugar source and nitrogen source may interact to form free radicals intermediates
during autoclaving (Maillard reaction)
・Phosphate and agar may interact while autoclaving to form H2O2 which can inhibit
the growth of some bacterial species and anaerobes.
・When autoclaving separately, remember the final total volume must still be 500 ml.
・I suggest (if autoclaving agar separately) to suspend 7.5 g of agar in 250-300 ml water.
・VERY DIFFICULT to fully dissolve 7.5 g agar in < 200 ml (please believe me!!)
・Use 200-250 ml ddH2O to suspend remaining materials in separate flask.
17
Preparing Materials for Lab
Tips for Pouring Plates
Allow media to cool to 55-50∘C in a water bath prior to pouring plates.
1. Open a new sleeve of sterile Petri dishes and take out 3-5 plates at a time.
・ Do not remove entire sleeve until done.
・ Plates are no longer sterile once removed from the bag.
・ Plates taken out of bag but not used must be sterilized by EtOH and UV before future use.
- I only sterilize/use loose Petri dishes in the rare occurrence we are totally out
(Including in the vault; will mention Petri dish sterilization later in PPT.)
2. Peel back the foil on flask and lift Petri dish lid just enough to pour media into bottom of plate.
・ Do not remove the entire foil cover on the flask.
・ Immediately re-cover the flask with foil when not actively pouring.
・ Avoid touching inner foil or rim of the flask.
- I also like to change my gloves prior to pouring plates to help minimize contamination risk
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Preparing Materials for Lab
Tips for Pouring Plates
Allow media to cool to 55-50∘C in a water bath prior to pouring plates.
3. Pour thin plates for lab exercises (Pour just enough agar to cover the bottom of Petri dish)
・ Thin layers of agar are okay for short-term incubation and storage, such as our lab exercises.
・ Prone to desiccation (drying out) so, plates must be used quickly if poured thin.
・ Thin plates will allow us to have more plates for the students too!
4. Pour thick plates for everything else (Pour about ½ volume of Petri dish)
・ Thick agar plates are needed for long-term incubation and storage.
- Such as for research purposes
・ If I ask for TSA plates to be made and do not specify it is for lab exercises, please pour thick :)
・Make sure not to pour so much that it touches the lid of the Petri dish, though! Toss those.
・Thick plates are much easier to streak on too!
19
Preparing Materials for Lab
Tips for Pouring Plates
Allow media to cool to 55-50∘C in a water bath prior to pouring plates.
5. Make sure the media is cooled to 55-50∘C prior to pouring!!
・ Extremely important, which is why it’s also included in each of the slides!
・ Plates poured too hot will result in excessive condensation.
・ Excessive condensation resulting in wet plates are unacceptable for several reasons:
- Can result in surface swarming by motile bacteria
- Causes smears that will prevent colony isolation
- Potential contamination from condensation on lid dripping onto agar
・ Condensation from plates CAN be evaporated by placing in 30∘C incubator OR
by leaving lid slightly open under biohood; however, this should not be necessary.
- Please be patient and allow media to cool down appropriately :)
20
Preparing Materials for Lab
Tips for Pouring Plates
6. When mixing autoclaved-separate media components:
・ Leave a stir bar in one of the flasks when autoclaving.
・ Allow flasks to cool in water bath to 55-60∘C.
- Letting it cool to 50∘C may cause it to start solidifying if you don’t work diligently!
- Mostly applies when mixing larger volumes (>100 ml) together
- This does not apply to heat-sensitive additives, such as antibiotics.
- Cool to 50∘C when adding things such as antibiotics!
・ Let flask stir slowly on hot plate (no heat) and slowly pour in the additional flask(s).
- This should preferably be done under a biohood.
- Transferring the contents of the other flasks by letting it run down the side of the flask
that’s stirring will help prevent bubbles from forming while mixing.
- Do not shake the flask to mix! You will get bubbles!
・ Bubbles can be popped by gently passing a flame over the surface of the agar.
21
Preparing Materials for Lab
Sterilizing Petri Dishes
Note: Petri dishes CANNOT be sterilized for use via autoclave!! They will melt/warp!
・ As previously mentioned, I only sterilize Petri dishes when we have absolutely NONE left in
our labs and the storage vault.
・ We tend to have plenty of loose Petri dishes from people removing EVERY plate from its
sleeve when pouring plates, but not using them all.
・Drives me crazy to see so many loose Petri dishes!!
・I normally set aside loose plates I find around to use for things like seed germination, etc..
To sterilize loose Petri dishes (done underneath biohood):
・Clean the inside of both the Petri dish and lid with 70% EtOH
・I like to spray thoroughly with EtOH and let it sit for a little before wiping with Kim wipe
・Leftover EtOH can evaporate during UV sterilization
・Turn on the UV light to sterilize by leaving the inside of the lid and Petri dish open
・Let Petri dishes sit under UV light for approximately 15 minutes.
22
Preparing Materials for Lab
Preparing Bacterial Cultures
1. Maintain Lab Bacterial Cultures by Restreaking onto TSA:
・Label pre-warmed TSA plates with the following:
・ Bacteria name
・ ATCC Number
・ Your initials
・ Date streaked
・ Media used
Pre-warming plates:
Incubate at 37℃, agar side down, for 15-20 min.
This decreases the ‘lag’ phase of bacterial growth.
・We normally start the semester with fresh ATCC cultures from KWIK-STIKS (Will show you guys how)
2. Streak Corresponding Bacteria with Sterile Loop:
・ Quadrant streak onto TSA
・ We need a minimum of four plates per each bacteria we’ll be using for the upcoming week
・ Given we have the TSA available, eight plates would be great so students don’t have to wait!
・ We incubate all our lab bacteria at 37℃
23
Preparing Materials for Lab
Culturing from Glycerol Stocks
1. Important: Do not let the glycerol stocks thaw!
・Think of it like ice cream! You can freeze ice cream after it’s melted a bit, but it’s never the same.
・Only take out a few glycerol stocks at a time and immediately transfer to ice block kept in -80∘C
2. We start the semester with fresh ATCC cultures from KWIK-STIKS (previously
mentioned), but if KWIK-STIKS aren’t available for each bacteria we use, culture from
glycerol stocks!
3. To culture from glycerol stock, lightly scrape the top of the surface with a sterile loop
and immediately transfer to TSA plate & incubate at 37∘C.
4. Return glycerol stocks to the same position in its box in the -80∘C freezer.
24
Preparing Materials for Lab
ATCC Bacterial Strains
Gram-Negative
・ Escherichia coli (25922)
・ Klebsiella aerogenes (13048)
・ Proteus vulgaris (49132)
・ Pseudomonas putida (31483)
Gram-Positive
・ Enterococcus faecium (8459)
・ Bacillus subtilis (6633)
・ Staphylococcus aureus (12598)
・ Staphylococcus epidermidis:
ATCC 12228: Biofilm-negative
ATCC 35984: Biofilm-positive
25
Preparing Materials for Lab
Controls for Lab Exercises
・ Phenol Red (Lactose)
E. coli (++)
P. putida (−)
・ MacConkey
E. coli (++)
K. aerogenes (+)
P. putida (−)
・ Simmons Citrate
K. aerogenes (+)
E. coli (−)
・ Mannitol Salt Agar (MSA)
S. aureus (+)
S. epidermidis (−)
・ SIM
P. vulgaris: H2S (+), Indole (+), Motile (+)
P. putida : H2S (−), Indole (−), Motile (+)
・ Starch Hydrolysis
B. subtilis (+)
E. coli (−)
・ Catalase
B. subtilis (+)
E. faecium (−)
・ Casein Hydrolysis
B. subtilis (+)
E. coli (−)
Students must use a
positive and negative
control for each lab
exercise!
26
Preparing Materials for Lab
Preparing Reagents
1. Aliquot Reagents from Stock Bottle into 50 ml tube(s):
・Pour underneath biohood and label with the following:
・ Reagent name
・ Reference number
・ Date poured
・ Your initials
Doesn’t apply to our smaller tubes
of expensive reagents, like for PCR!!
More on that in a later slide!
2. Aliquot Reagents from 50 ml tube(s) into 5 ml tubes:
・ Depending on the reagent, this won’t have to be done under the biohood (ex. 0.1x Crystal Violet)
・ Considering there’s 16 students per lab section, I try to have 8 tubes per lab section.
・ Keep an extra 50 ml tube with reagent just in case (so we don’t have to run back and forth.)
・ We can use 15 ml tubes if 5 ml tubes are unavailable.
*Note: Broths must be poured underneath the biohood! Don’t open stock broths outside of the hood! 27
Preparing Materials for Lab
Preparing 1x PBS
Preparing 1x PBS from 20x Stock Bottle:
・ 475 ml of ddH2O and 25 ml of 20x PBS into 500 ml bottle
・ Invert several times to mix
・ Autoclave as per standard protocol
PBS contamination seems to be a problem as of late, so…
Phosphate Buffered Saline (PBS) is
the buffer we most commonly use
for bacterial suspensions or serial
dilutions. Must keep in stock!
Unlike water, PBS helps prevent
cells from rupturing or shriveling
due to osmosis while in
suspension.
・ After allowing autoclaved PBS to cool, transfer some to a 50 ml conical tube under biohood
・ Spot 10 µl of PBS onto TSA (3-4 times) and allow to drip down plate (as per Track Dilution)
・ Incubate TSA plate at 37∘C for 24 hours
・ Check that there’s no growth prior to using for exercises or projects
28
Preparing Materials for Lab
Tips for Preparing Chemicals by Molarity
IMPORTANT! Bottles of powdered chemicals will state the formula (FW) or molecular weight
(MW) of the compound on the front. This is NOT the molar mass (g/mol)!
・If you cannot find g/mol on the bottle, make sure to look it up prior to using.
・Refresher: Molar mass (g/mol) = # of grams needed to make a 1 M solution in 1 L of solvent
・We normally make chemical stocks in 100 ml bottles (So, 1/10th of a liter)
・To quickly make a 100 ml (1 M) stock solution, just move the decimal in the molar mass to the left!
Ex. NaCl molar mass = 58.44 g/mol
・58.44 g into 1 L of water ➝ 1 M NaCl solution
・5.844 g into 100 ml of water ➝ 1 M NaCl solution
・Need to make a 100 ml stock of 100 mM solution? Just move that decimal over to the left again!
・5.844 g into 100 ml of water ➝ 1 M NaCl solution
・0.584 g into 100 ml of water ➝ 100 mM NaCl solution
29
Preparing Materials for Lab
Preparing Reagents for MALDI-TOF MS
Reagent Locations:
・Stock bottles of ethanol and water are kept in Dr. LaMontagne’s lab (B3104)
・Must use HPLC EtOH and LC-MS Water! ‘For mass spectrometry’ indicated on the bottles
・Stock bottles of Formic Acid and Acetonitrile are in the yellow flammable cabinet (autoclave lab)
・Working solution of 80% TFA and the Bruker Standard Solvent in the gray corrosive cabinet in
autoclave lab
・80% TFA used for cleaning steel targets; Let Dr. LaMontagne prepare the working solution if out
・ONLY work with TFA, Acetonitrile, and (concentrated) Formic Acid underneath a fume hood!!
・Bruker Standard Solvent used for preparing Matrix
I believe we recently started using a new type of Matrix, though!
・BTS is kept in the -20∘C freezer in the autoclave lab
Please help keep track of our
MALDI-TOF stocks!
If we get low, we need to order
more ASAP!
30
Preparing Materials for Lab
Preparing Reagents for MALDI-TOF MS
Preparing 70% Formic Acid Stock from 100% Stock Bottle
・ Begin by rinsing out a 100 ml bottle with leftover 70% Formic Acid
Prepare the following under a fume hood:
・ Add 30 ml of LC-MS grade water to 100 ml bottle
ALWAYS ADD ACID TO
WATER!!
・ Add 70 ml of 100% Formic acid
NOT WATER TO ACID!!
・ Screw bottle closed and invert to mix
100 ml is enough
for 2,000 preps!!
Remaining reagents for MALDI-TOF (100% Ethanol, Water, 100% Acetonitrile…) can all
be prepared by transferring to a 50 ml conical tube:
・ Prepare Ethanol and Water underneath biohood
・ Prepare 100% Acetonitrile underneath fume hood
31
Preparing Materials for Lab
Preparing PCR Reagents
16S PCR reagent volumes per 25 µl reaction:
・Platiunum II Master Mix
・Nuclease-Free Water
・Forward Primer (10 µM)
・Reverse Primer (10 µM)
・DNA template
12 µl
10 µl
1 µl
1 µl
1 µl
Our stock primers come in
concentrations of 100 µM. So, if you
need to dilute for PCR:
Make 100 µl stocks of 10 µM primers
using 90 µl of PCR-Grade Water and
10 µl of primer stock.
When preparing reagents for students to use in lab:
・Thaw frozen reagents on ice (Platinum II and Nuclease-Free Water)
・Transfer enough for each student (and a little extra for demonstrations/cushion) to separate 1.5 ml tubes
・16 students per lab section, so I would suggest transferring enough for 20 reactions:
Tube 1: Platinum II (240 µl)
Tube 2: Nuclease-Free Water (200 µl)
Tube 3: Forward Primer (20 µl)
Tube 4: Reverse Primer (20 µl)
Make sure to spin down
reagents prior to using!!
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Preparing Materials for Lab
Waste & Biohazard Disposal
1. Uncontaminated nutrient broths can be disposed of via sink
・ No risk of solidifying (no agar) and does not contain chemicals toxic to the environment.
2. Contaminated broths must be autoclaved prior to disposal
・ In the event you have a large volume of broth (> 500 ml) and a large amount of biomass…
- I suggest draining liquid into sink after autoclaving, and then dispose of biomass in biohazard bag
3. Agar can be tossed into a biohazard bag for autoclaving & disposal
・ Pour into biohazard bag and double bag
・ Tape closed with small opening for ventilation
- Sealing the bags too tightly will cause them to pop in the autoclave!
・ Nothing with agar or agarose is allowed down the sink!!!
・ When cleaning flasks used to make agar: Soak in soap to breakdown any extra agar!
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Preparing Materials for Lab
Waste & Biohazard Disposal
4. Dispose of chemicals in a chemical waste container (No chemicals in sink!)
・ We keep a chemical waste container under the fume hood in the autoclave lab
・ Gram-staining reagents are disposed of in a bottle in the micro lab
5. If Petri dishes are not parafilmed, tape the sides of a stack with tape before bagging
・Loose/open lids can cause the agar to seep out (that gross black gunk you see in the bins)
・Mostly an issue if the bag were to pop, but still best to avoid
6. Glass culture tubes must be autoclaved and disposed of via Broken Glass Container
・ Autoclave the culture tubes as you would when making culture tubes
・ Remove the caps after autoclaving (we keep those!)
・ Let solid media tubes solidify before tossing into broken glass container (to minimize mess!)
・ Culture tubes with broth, namely unused broth, can be poured down sink prior to tossing tube
into broken waste container
・ Try to just place the glass tubes in the container… No need to throw and make them all shatter!
34
Microbiology Lab (B3211) Map
(I’m proud of this so including it heh)
Hallway
SAFETY
SHOWER
Autoclave
Lab (3213)
Desk
Bench
Counter
Sink
FIRST AID KIT
Fume
Hood
BROKEN
GLASS
DISPOSAL
REGULAR
TRASH
EYE
WASH
Lab
Bench
Lab
Bench
BIOHAZARD
WASTE
BIOHAZARD
WASTE
Cabinet
REGULAR
TRASH
Hallway
GRAM-STAIN
WASTE
Sink
Counter
35
GLOVES
Preparing Materials for Lab
Setting up for Lab:
Before Lab
− Prepare media at least two days before lab
・ To make sure plates aren’t too wet with condensation (no wet plates!!)
・ Will also help verify no contamination was present in the media prior to pouring
− Have reagents prepared and aliquoted day before lab
・ Unless a protocol calls for something freshly prepared!
− If possible, set aside materials we’ll need the day/evening before lab
・ Make sure to label them very obviously that they’re for lab exercises (so no touchy-touchy)
・ Doesn’t apply to materials that everyone uses regularly, like pipettes!!
・ However, we need to make sure we have a count of the number of available pipettes/track
down the others prior to lab if need be
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Preparing Materials for Lab
Before Lab
Test out the exercises we will be doing prior to lab
・ To make sure everything is working properly!
− Preferably 48-72 hours before lab if exercise requires culturing (Monday is best)
・ This will allow us to have time to figure out what went wrong with the exercise
(should it not work) and prepare fresh materials for the lab.
− Make sure you understand the concept(s) of the exercises too
・ Aside from it just being good to know..
・ Understanding concepts will help you troubleshoot any potential problems.
・ You’ll also be able to better answer questions from students :)
Please send me photos of your results, if possible, for my own peace of mind :)
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Assisting
During Lab
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Assisting During Lab
Lab Flow
− Labs will start with brief explanation of exercises along with demonstration(s)
・ Be mindful of students who come in late: Dr. LaMontagne and I won’t repeat
ourselves for them/show another demonstration, but we need to make sure they’re
doing the exercises properly!
− Students will then begin exercises based on which bench they’re at…
・ Due to the lack of pipettes available per student, I normally let eight students at one
bench begin pipette exercises while the eight students at the other bench begin
culturing or microscope exercises, etc.
・ Additional materials for each exercise will be on the back bench; students will grab
and replace materials as they go along in lab.
➝ If you see a student hoarding materials/equipment while not using them,
tell them to put it back so others can use them!
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Assisting During Lab
Lab Flow
− Please keep an eye on whether materials are getting low…
・If you see students are about to run out of certain materials, like reagents, pipette
tips, etc… Please go ahead and refill/replace them so we can keep the lab moving!
− For culturing exercises, we’ll have the students stack their plates on the front bench
・Please make sure that all Petri dishes are upright (Lid down, agar up) when storing in
the metal canisters for incubation!!
・Label canisters with the exercise name, date, and the lab section (W9, W1, T9, or F9)
➝ If it’s a culture tube exercise or something we’re storing for future use (DNA, etc.)
please label the racks as well!!
・Once each student is done, incubate the canisters and/or store tubes in proper place.
➝ Bacteria for every lab exercise we do (with ATCC strains) are incubated at 37∘C
➝ EtOH preps in fridge (4∘C) and FA extractions/DNA in freezer (-20∘C)
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Assisting During Lab
Supervising Students in Lab
− Make sure students are following safety guidelines while in lab, including…
・ Wearing proper PPE (Lab coat, closed-toe shoes, goggles/glasses, gloves)
・With the exception of using the Bunsen burner… No gloves and flames!
・I sometimes fall short in enforcing the use of goggles; NOT because I don’t care,
but because it will slip my mind. We must enforce PPE and I will be more mindful, but…
・Please help me enforce the use of goggles, especially when we use reagents like
in Gram-staining, etc.
・ Using proper aseptic techniques (Cleaning bench before AND after, etc.)
・ Using equipment properly
・ Leaving any food or drink outside (includes bottled drinks - nothing consumable allowed!)
Because students are learning (and many have little lab experience), it’s common for mistakes to be made.
However, students blatantly disregarding safety guidelines will be removed from the lab.
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Assisting During Lab
Supervising Students in Lab
− Students can converse but make sure they stay on track!
・Should go without saying but EVERY student must do the lab exercise(s) themselves!
・They can talk amongst each other (it weirds me out when they’re all silent), but
they need to stay focused and complete their lab exercises, alone, in a timely manner.
・However, talking ≠ goofing off/disruptive behavior.
・If students have lab exercise questions, they should ask you or me, NOT each other.
➝ They’ll just confuse each other!
− Things students seem to often do… (so, watch out for!)
・Put P1000 tips onto P200 pipettes by forcing it over the piston which is a NO-NO.
・Trying to use a pipette outside of its’ intended range, another NO-NO.
・Leaving their Bunsen burner on when not in use – Tell ‘em to shut it off!
・Leaving Petri dishes open when not actively streaking.
・Labelling the LID of the Petri dish, rather than the agar-side (Correct them if you see this!)
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Assisting During Lab
Lab Books
− Near the end of each week’s lab, Dr. LaMontagne and I will grade students’ exercise
entries and leave feedback.
・ Lab exercises that have same-day results (i.e., don’t require culturing) can
be graded the day it’s performed/before they leave.
・ Exercises that DO require culturing (so, waiting for results) will be checked/graded
the following week
− After having their books graded, they’ll show you their book and you’ll enter in their grade.
・ I’ll provide an excel spreadsheet with separate sheets for each lab section.
・ Make sure their lab book pages have either my or Dr. LaMontagne’s initials AND a
grade on it.
- I HAVE had students try to BS lab book grades by ‘grading’ it themselves.
- Which is cheating, obviously. And just so stupid. That was back in Fall 2020.
- I gave those students a 0 for the week and informed Dr. LaMontagne.
- If this happens, let me or Dr. LaMontagne know immediately.
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Assisting During Lab
Lab Books
− You will be asked many, many times by students… “Does this need to be in my lab book?”
・The answer is YES! ALWAYS YES!
・EVERYTHING done in lab (exercises, project, etc..) needs to be in their lab books!
− Remind students that it’s ONE exercise per page if you catch them cramming it all together!
− Other questions you will definitely be asked:
・Can I write on a separate sheet and transfer it later? ➝ NO!
・Recording research/lab work MUST be done while they are working!
・(Don’t let them stop in between each steps to record in their books though…)
・If they forget their lab book, they can write on a separate sheet and either TAPE or STAPLE in
➝ NO REWRITING OR TRANSFERING!!
・Can I write in pencil? ➝ NO!
・They’ll go on about how they want it pretty, yadda yadda, but work must be recorded in PEN!
・What do I do for the ‘Next’!?!? ➝ Use their critical thinking!
・For me, there’s not really a “wrong” answer if I can see where they’re coming from/it makes logical sense
・However, it’s NEVER EVER “to clean up” or just whatever it is they’re actually do next in lab.
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Please review lab exercise PPTs for more detailed info :)
Media Used in
Lab Exercises
The following section includes…
・ Purpose of each media
・ Media composition
・ Interpretation of results
・ Bacterial controls needed
(Lots of info crammed into as few slides as possible; little messy!)
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(Week 6)
SIM (Sulfur, Indole, Motility) Agar
SIM agar is a differential medium that tests the ability of an organism to do several things:
1)
2)
3)
Reduce sulfur to hydrogen sulfide (H2S)
Produce indole (Through production of tryptophanase, which can convert tryptophan to indole)
Move through agar (motile)
Medium contains:
・ Casein and Meat Peptones (Nutrients)
- Supplies AAs, like tryptophan, and nitrogenous compounds
・ Sodium thiosulfate
- Acts as the substrate for enzymatic sulfur reduction
・ Ferric ammonium citrate (Indicator)
- Reacts with H2S to form ferrous sulfide, a black precipitate
・ 0.35% Agar (Semi-solid for motility)
- Semi-solid ≠ Broth (No liquid-y tubes!)
Note: After incubation, five drops of Kovac’s Reagent is required to
visualize indole production (If positive, red ring will form on top)
Positive Control: P. vulgaris
Negative Control: P. putida (+ for motility)
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(Week 7)
MacConkey Agar
MacConkey agar is a selective and differential medium that tests for lactose fermentation
Medium contains:
・ Lactose (Fermentable carbohydrate source)
· Encourages the growth of fecal coliforms and provide a means of
differentiating them
・ Bile Salts and Crystal Violet (Selective reagents)
· Serve to encourage Gram-negative growth by inhibiting the growth of
most Gram-positive organisms
・ Neutral Red (pH indicator)
· Turns red at a pH below 6.8 and is colorless at any pH greater than 6.8
Uninoculated MacConkey Agar
MacConkey also assists in visual distinction of E. coli from
other lactose-fermenting enteric Gram-negative rods
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(Week 7)
MacConkey Interpretation
Fermentation Positive Results:
· Strong fermenters - Colonies will appear pink and be
surrounded by zone of acid precipitated bile (Ex. E. coli)
· Fermenters – Colonies will appear red or pink
Fermentation Negative Results:
· Colorless colonies indicate that the bacteria does not
ferment lactose and is not a fecal coliform
· Some lactose fermentation-negative bacteria can also increase
the pH of the media, resulting in a peach/yellow color
· The pH indicator (Neutral red) becomes colorless at pH > 6.8
(Agar is naturally a tan-ish color)
Strong Positive Control: E. coli
Positive Control: K. aerogenes
Negative control: P. putida
I want students to use all three
as seen in the photo attached
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(Week 7)
Mannitol Salt Agar (MSA)
Mannitol Salt Agar (MSA) is a selective and differential medium that tests for mannitol
fermentation and differentiates pathogenic Staphylococci
Medium contains:
・ Mannitol (Fermentable carbohydrate source)
· A species signature of Staphylococcus aureus
· Discriminates it from most other members of the genus
・ 7.5% NaCl (High Salt Concentration)
· Serve to encourage Staphylococci growth by inhibiting the growth of
most other Gram-positive and Gram-negative organisms (Not all!)
・ Phenol Red (pH indicator)
· Red at a pH > 7.4 and turns yellow at pH < 6.8
MSA allows for also visual differentiation of S. aureus from other
Staphylococcus spp. based on color change from red to yellow
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(Week 7)
MSA Interpretation
· Able to grow in 7.5% NaCl
· Able to grow in 7.5% NaCl
· Mannitol fermentation-positive
colonies will result in pH decrease,
turning media yellow
· Mannitol fermentation-negative
colonies will appear red/colorless
· Unable to grow in 7.5% NaCl (Growth
inhibited by high salinity)
· Mannitol fermentation can’t be
assessed (N/A) without growth!
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Simmons Citrate
(Week 7)
Differentiates coliforms from fecal coliforms based on the ability of non-fecal coliforms to
grow with:
・Citrate as the sole carbon source
・Ammonium as the sole nitrogen source
Students should include something
about this in their goal AND conclusion!
Medium contains:
・ Citrate
・ Ammonium ions
・ A pH indicator (Bromothymol blue)
When the bacteria metabolize citrate, the ammonium salts are broken down to
ammonia, which increases pH (Growth raises the pH)
Bromothymol blue is green at pH < 6.9, and then turns blue at pH > 7.6
- Make sure to autoclave these in a tilting rack!
- After autoclaving, tilt the tube rack onto its side to allow the
media to solidify at a slant
Uninoculated Simmons Citrate
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(Week 7)
Simmons Citrate Interpretation
Positive Results:
Blue color change - Indicates the bacteria is able to use
citrate as its sole carbon source and ammonium as its sole
nitrogen source
Ex. K. aerogenes (Non-fecal coliform)
Negative Results:
No color change (medium remains green) - Indicates the
bacteria is unable to use citrate as its sole carbon source and
ammonium as its sole nitrogen source
Ex. E. coli (Fecal coliform)
K. aerogenes
E. coli
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(Week 7)
Phenol Red for Lactose Fermentation
Tests the ability of a bacteria to ferment carbohydrates (Lactose for us)
Medium contains:
・ A carbohydrate source (lactose, glucose, etc.)
- Add 5 g of Lactose per 500 mL when preparing
・ A pH indicator (phenol red)
・ Durham tube (for capturing gas)
- Add Durham tube to each tube after adding broth but before autoclaving.
- Pipetting some broth into Durham tube will help prevent bubbles.
(Will likely go away after autoclaving; if it doesn’t, bubble will expand over time)
The production of acid lowers the pH of the medium, which is detected by the color
change of the pH indicator. Gas production is indicated by the presence of a bubble in
the Durham tube. Phenol red is yellow at a pH < 6.8 and red at a pH of > 7.4
・ If lactose is utilized, the pH indicator phenol red will turn yellow (acid production) or pink (alkalinization/ammonia production)
・ If gas is produced, it will replace liquid in the Durham tube
Positive Control: E. coli (Acid and Gas production)
Negative Control: P. putida
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(Week 8)
Skim Milk Agar for Casein Hydrolysis
Skim milk agar is a differential medium that tests for the ability to hydrolyze casein through the production
of the exoenzyme caseinase (Students must include this in goal)
Medium contains:
・ Glucose (Carbon source)
・ Yeast extract (Provides vitamin B complex)
・ Skim Milk (Casein source; cow milk has ~ 80% casein)
・ Casein enzymic hydrolysate (Provides AAs & nitrogenous compounds)
PLEASE REFER TO MEDIA TIP #4 (Slide 13)
WHEN MAKING SKIM MILK AGAR!!!
Following 24 hour incubation at 37∘C, results can be analyzed by the presence or absence of
a zone of clearance surrounding the bacterial growth. Zone of clearing is indicative of
caseinase production resulting in casein hydrolysis. No zone of clearing indicates the
bacteria was unable to produce caseinase to breakdown casein the in media.
Positive Control: B. subtilis
Negative Control: E. coli
E. coli
B. subtilis
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(Week 8)
Starch Agar for Starch Hydrolysis
Starch agar is a differential medium that tests for the ability to hydrolyze starch through the
production of the exoenzyme amylase (Students must include this in goal)
Medium contains:
・ Beef Extract
· Provides the nitrogen, vitamins, carbon and amino acids
・ Soluble Starch
· Makes up approximately 40% of the media; used to assess
amylase production
Following 24 hour incubation at 37∘C, the addition of Gram’s Iodine is
required to analyze amylase production.
・ Gram’s Iodine binds to starch molecules, resulting in a dark blueblack color. If no starch is present for iodine to bind to, a zone of
clearance will be present.
Positive Control: B. subtilis
Negative Control: E. coli
E. coli
B. subtilis
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Lab Practicals
The following section includes…
・ Practical 1: Track Dilution
・ Practical 2: Unknown ID Flow Chart & Info
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Practical I: Track Dilution
Practical Overview
・ Students will prepare an E. coli suspension and measure the A600
・ Using the A600, students will dilute their suspension to 105 CFU ml-1
・ Students will then further dilute their 105 suspension to 103 CFU ml-1
・ Students will spot four 10 µl spots onto the top of a TSA plate (2 spots of 105, 2 spots of 103)
・ Plates will be incubated at 37∘C for 24 hours
・ Grade will be based on whether they get isolated colonies after 24 hours.
Isolated colonies = 5 points
Please refer to “CFUs and Track Dilution” PPT for more detailed info :)
Students may use a calculator for their dilution math, but they must work alone!
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Practical II
Start Here
Gram-Stain
Gram-positive (+)
Gram-negative (−)
Catalase Test
(−)
E. faecium
(+)
Starch Hydrolysis
S. epidermidis
(−)
(+)
Mannitol Salt Agar
(MSA)
(−)
(−)
MacConkey Agar
(Lactose fermentation)
(+)
(+)
S. aureus
SIM
Simmons Citrate
K. aerogenes
(−)
E. coli
(+)
P. vulgaris
(−)
P. putida
(+)
B. subtilis
Unknown ID Practical Flow Chart
Each test/assay must be on its own page!!
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Practical II
Unknown ID: Codes & Info
Letter Code(s)
Bacteria
Gram-Stain
Biochemical Assay Results
A, I
K. aerogenes
Negative
MacConkey (+), Citrate (+)
B, J
B. subtilis
Positive
Catalase (+), MSA (-), Starch Hydrolysis (+)
C, K
S. aureus
Positive
Catalase (+), MSA (+)
D, L
E. coli
Negative
MacConkey (+), Citrate (-)
E, M
E. faecium
Positive
Catalase (-)
F, N
P. putida
Negative
MacConkey (-), SIM (+ motility only)
G, O
S. epidermidis
Positive
Catalase (+), MSA (-), Starch Hydrolysis (-)
H, P
P. vulgaris
Negative
MacConkey (-), SIM (+ for each)
・ Label TSA plates with just a letter and the date, and streak the corresponding bacteria
・Need 16 plates per lab section/day ➝ One stack of 16 plates will be okay for both Wednesday sections
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Some Additional
Comments
and Suggestions :)
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Microbiology Lab TAing
Most Important Rules for Micro TAing (to me, at least):
1) Dr. LaMontagne comes first
2) Dr. LaMontagne knows best
・I do my best to make the labs as easy/stress-free for Dr. LaMontagne as possible.
・We need to make sure he has all of the materials he needs ready to go so he can come in and teach.
・If Dr. LaMontagne asks you to do something while you’re TAing for my lab section(s), you go do it!!
・Dr. LaMontagne can use any materials he needs and throw away anything he sees fit.
➝ I often label things I need as “don’t use” which applies to everyone but Dr. LaMontagne
・I’ve always tried to act as a “buffer” between him and repetitive/dumb questions from students. (There’s a LOT!)
I’d prefer them to blow up my email vs. unnecessarily blowing up his!!
➝ I’ll have you guys help me with handling student emails too
・If I do something that differs from what Dr. LaMontagne explicitly stated to do, please let me know!
・This should go without saying, but Dr. LaMontagne really does know best. If he tells you to do/prepare
something in a certain way, there is a reason for it and you need to do it.
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Microbiology Lab TAing
Things to Remember:
・You ARE going to make mistakes from time to time
・Sometimes things don’t turn out properly or work as they should
・AND THAT’S OKAY!
・Do NOT beat yourself up over it!
・‘Mistakes’ are a great learning tool if you choose to see it as such.
…Hopefully they’re not hazardous/expensive mistakes though :)
・There’s a difference between making a mistake and being ‘lazy’ with shortcuts, etc.
・Sloppy work is NOT a mistake! That’s a choice!
・(I know I won’t have to worry about that with y’all!)
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Things That Helped Me TA!
Things that help make TAing/instructing micro lab FUN for me!
・Having a deep, genuine interest in microbiology and research
This slide is more so
on mindset :)
・Seeing (and appreciating) TAing as an opportunity
・Everyone’s different, of course, but I personally never thought of TAing as a “job”
(in the sense of ‘I’m getting paid so I have to do it’ yadda yadda)
・I always have (and still do) see TAing/instructing as an opportunity to improve my
knowledge of microbiology, my skills in lab techniques, and conveying research/science to others
・An opportunity you get paid for, which only makes it better! (It IS a job though, don’t forget that!)
・Wanting to help Dr. LaMontagne
・My admiration and appreciation for Dr. LaMontagne is well known, but having a ’boss’ that inspires you
to do your best is few and far between.
・I appreciate getting to assist him in lab and do my best to make lab easy for him (hence previous slide)
➝ I want everyone to help make lab easy for him too!!
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I hope this is helpful
to you guys! :)
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