WW and General Lab Skills, Tests, and Techniques

sandwich elisa

antigen is sandwiched between a capture Ab (which grabbed it) stuck to the plate and a detection Ab bound to some kind of indicator that is administered to the plate after the antigen

most commonly used as it is both sensitive and specific

single plex

detects a single target antigen

reading an elisa

the intensity of the color indicator is directly proportional to the concentration of the target antigen in the sample

usually have 2 replicates per curve and per sample dilution

no bubbles in the sample wells - messes with the reading

if you suspect the signal will be low, you may want to use a pipette over a plate washer for the washes, as the pipette is more gentle

finished plate should be read immediately

anti-target Ab-reporter enzyme complex - the reporter enzyme works on the bound substrate to produce a colored signal when the Ab binds the target

may add a plate blocker (irrelevant protein that blocks unsaturated bound Ab sites) to prevent miscounting - binds up all remaining free binding sites on capture antibodies after sample has done its thing

elisa tricks

clingfilm - if you have nothing else, you can cover a plate with cling film to keep it from drying out

low signal - if you aren’t seeing much, you can incubate longer

vortex Abs prior to use

polyclonal Abs are often used as capture Abs as they can be made to be specific to multiple epitopes on the same target antigen to increase capture rate

after washing an elisa

you’re going to want to put some paper towels down on the bench top then invert and slam down that plate several times

be aggressive about it

EIA

another name for an elisa

indirect detection

greater elisa sensitivity because multiple signal-bound Abs can bind the primary Ab with the target, amplifying the signal and potentially turning the indetectable detectable

alternative - use molecules that can bind up multiple biotin (a signal option) like avadin to further increase signal - so detector Ab hosts multiple signal molecules via an intermediary protein to increase signal strength

biotinylated Abs bind avidin or streptavidin, which bind multiple signal molecules

cell culture

keep culture vials upright when handling

transfer flask from incubator to hood quickly to avoid gathering condensation, which makes it hard to see the monolayer inside when you are trying to get the cells off

loosely adherent cells just need fluid run over the monolayer to ger them to detach - strongly adherent cells need the addition of an enzyme like trypsin

prepare new culture flasks before passaging

careful not the tip the fluid into the filter in the cap

don’t scrape pipette tip on the monolayer

replacing the media every 3 ish days is a rule of thumb

60% confluence is when you can almost double your cells and have space to fit all of them in

make sure the whole bottom of the flask is covered in media - no tension holes

culture split factors are written as proportions, with the first number referring to the original amount and the second referring to the new amount to be grown

e.g. if using all the same sized flasks, a 1:2 split means taking your cells from the first flask and splitting them between 2 new flasks

increasing or decreasing the size of the new flask relative to the old changes the ratio - larger relative flask means more space for more cells to grow

passaging cells

1. prepare your new flasks

2. aspirate media from the old flasks and check the monolayer

3. Aspirate 10 ml PBS and run it over the monolayer, repeatedly aspirating and releasing it

4. can cap the flask and hold it up to the light to check if there’s any of the cells still adhering

5. Once the majority of cells are suspended, aspirate up the PBS cell suspension and deposit into a tube

6. Spin down

7. Aspirate PBS and resuspend the cell pellet in media, mix cells to resuspend

media

if you need to use media from a commonly used bottle, it is best practice to pour some into the reservoir to pipette from rather than going directly from the bottle

pipetting and serologicals

wet the tip by aspirating a little, completely depressing the plunger, then taking in the measured amount for increased accuracy

you can use a serological to suck bubbles and foam off the top of a liquid

if transferring a very drippy liquid, you can carefully tip the bottle opening close to where the liquid is going to go to reduce the impact of dripping

really jam the multichannel tips on there

the smaller an amount, the harder it is to measure it accurately, so you can sometimes dilute something to be added (factor of 10 usually) so you can add a larger amount for increased accuracy

fluid in a pipette tip will usually stay put, so you can tilt the pipette and stuff

if you have very little fluid to spare, a single pipette is better over a multichannel

ALWAYS watch the level of what you are aspirating - don’t suck up to much fluid and get it into the filter

if possible, mixing cells with a p100 over a serological can be faster and create less bubbles

depress pipette to the first resistant before inserting into fluid to take it up to make sure you don’t get any air in there

if pipetting something very viscous, and you aren't working with sterile, you can use scissors to diagonally chop the top to increase the side and speed of draw. Also, depress pump a bit below first resistance when aspirating, but depress only to first resistance when depositing.

middle channels in a multi pipette are more reliable

have tip make contact with the side of a well if possible before depositing to get everything out

when you need to add a precise amount of solution using a serological (say 13.1), seros aren’t very accurate at that level

best to either round up to the next whole number, add that, then use a pipette to remove the excess, or round down to the next whole number and use a p1000 to get the rest

aseptic technique

place bottles as far away as possible while they are within reach to avoid waving your sleeve over them

place open caps to the right

if something has autoclave tape on it, it is sterile and should only be opened in a hood

if you can, its often better to use serologicals to transfer fluids between vessels, even if you aren’t working under aseptic conditions, it’s still usually better than pouring

if you have something you want to make sure stays contained, you can put parafilm over the cap as an extra layer of protection

spray down cell cryovials heavily

spray hands before entering the hood

only allow the replaceable tip of the pipette to enter a flask

open seros like bananas

other

you can use the brown backing from the metal plate stickers to smooth it over the wells

you can leave a cell pellet dry for 4-5 minutes at most

wrap a kim wipe around an ampule before you break it - all parts go into sharps

when thawing a cryovial, set a timer but also check on it - when only a small core of ice remains, start bringing it over to your bench - the ambient heat from holding it should melt the rest so it is ready just as you need it

2 min timer to thaw small cryovials

get the media prepped in tubes for the cells first BEFORE thawing them

when mixing cells, its good practice to safely invert the tube at least once

cleaning

spray kim wipe rather than inside the hood directly to keep ethanol off the more delicate equipment

new gloves if moving from wider lab to TC

can raw dog a bleach solution by using a falcon tube to measure out roughly a 10th of what you’ll be needing

media can stain the hood - wipe up with di water before applying ethanol

to get rid of a full bin of sharps, close the red bin with the black lid (should click into place), then bring to the room on the far left by the elevator and zip tie it shut through the two holes, then put a red sticker on it. For the biohazard bins, wheel them over to the same room, take off the black plastic lid, tie up the first bag, (put two corners away, fold in the middle of the bag one over another, then double knot shut), then tie up the second bag the same way, then fold up the box and tape the flaps shut along the seam, then tape perpendicular, then two more strips parallel to the first over the seam, then put red sticker in company square on side of the box. To make up a new biohazard box, unfold a box upsidedown, tape up the bottom the same way you tape up a full box, flip over, double bag, put back in the coaster bottom, and be on your way.

when mixing water and bleach, bleach goes first

when cleaning the hood

1. spray kim wipes and wipe down hood and anything you used

2. plug in pipette boy

3. deposit seros in sharps

4. close hood

5. UV button

storage and organization

if you need to put a new rack in a freezer, especially a nitrogen freezer, you have to put it in there empty first to chill down to temp before you put samples in it

140 proof ethanol is 70% - 200 proof is 100%

xylene will melt plastic if left for a while, so it has to be stored in glass long term

label all media and solutions with the components and amounts/conc/percentages of each as well as your initials and the date it was produced

cell counting

(2uL AO + 18 uL cell suspension)/2 = 10 uL mix inserted into counting slide

to calculate the cell you need in your flask:

1. multiply desired density of flask area by the total number of cells required

2. divide product by the density of the cells you have (from the count) to get the cells you need in mL

3. mult by 1000 to get it in uL

4. substract this number from the total volume of fluid allowed in the flask to get how much media you need

5. Add the right amount of media, pipette in your cells, mix via aspiration, and done

C1V1 math

(original conc of stock)(how much of it you need to add to the solution) = (new concentration you want of it)(final solution volume)

C has to be in moles, so you’ll have to convert

the m or u or general unit has to be consistent throughout

C1V1 alternative

volume I need to add = (conc percentage needed)(solution volume)

dalton

atomic mass unit - used to measure protein size

1Da is about 1g/mol

molecular weight of a protein typically given in KDa

Molarity

M = moles

X = solution conc

W = molecular weight

moles of solute (quanitity) per liter of solution

M = (X * mol)/(W * L)

mM = (conc in mg/mL)/(W * kDa)

w/ 2mg/mL and 79.4kDa

= 0.025mM - 25uM

IN SUM:

(mg/mL)/(W in kDa) = mM

(ug/uL)/(W in kDa) = uM

dilutions

for a 1:X dilution, just divide the final desired volume with that X to get the factor of your dilution component

should be 1:10 dilution is 1 part conc to 9 parts diluent for a total of 10 parts

stains

DAPI - nucleus

AO - vesicles - death

flow cytometry

first chart is usually SSC (y) by FSC (x)

bottom left quadrant is neither, top right is both

FSC = cell size

SSC = cell complexity (how granular)

dead cells tend to fluoresce more - another way of telling them apart beyond size and complexity

doublets tend to have a larger area but the same height as single cells, and thus can be screened for that

useful to include positive and negative cell markers - markers on your cells of interest and markers only on other cell types

usually have to block Fc receptors otherwise they can generate false positives

have to use an FC blocker to prevent off-target fluorophore binding before staining with markers

have to wash after all staining steps

need to leave cells in formalin overnight to make them permeable to intracellular markers - do this after extracellular staining since this process can mess with their binding sites

cell markers

CD3 - T cells

CD4 and CD8 - helper and killer

H and E staining

the timing for the washes can be a bit lax, but you MUST be precise for the timing of the staining, ESPECIALLY THE BLUING

for the time sensitive staining, make sure the caps for the di water and waste are open so you can immediately transfer the slide and such

you can stick a still-wrapped serological on the pipette boy to quicky unsheath it when needed

keep the slide wet - go from staining steps to water bath steps

can use kim wipe to dab around the tissue slice (NEVER ON) to remove excess liquid

use p 1000 to gently drip fluid onto tissue

poke cover slip in resin with plastic pipette to work out bubbles

3 is a manageable number to handle at one time

be careful not to let the slides stick to each other

you can leave the slides in water for longer than specified if you need some more time to handle the others - does NOT work with xylene tho

you can bulk stain the slides but NOT on the bluing step since its so time sensitive

best to get multiple water baths for the last bath to keep it to 2 slides per bath to minimize sticking risk

microtome

soak the blocks in ice water (not warm water - screws the wax) for like 10 minutes before slicing for less fraying - may want to trim the block a bit to make a fresh edge then soak then slice

softer wax can prevent slice curling

only clean the blade parallel to the edge - keep as sharp as possible

1. need a di water bath and di water to soak the blocks

2. spray down the tub with ethanol prior to filling to remove dust

3. put beaker with wet ice and cold water shover into a larger ice bucket to keep the bath chilled

4. use flat paintbrush to pat the inserted blade into place

5. give blocks a cursory pat dry before loading

6. trim block surface smooth before going to slice

7. skim the bath surface with a kim wipe to remove any floating debris before putting in the slices

8. troubleshooting methods include - changing the blade, changing blade types, using a different part of the blade, changing blade angle, warming the wax a bit, rehydrating the block

9. Use brush to gently scoop up the slices to transfer them to the bath

10. tissue goes on the textured side of the slide

11. Dip the glass slide underneath the desired slice and gently pull up at an angle to capture

to clean the microtome

1. slides have to be incubated overnight

2. remove blade then dust off wax

3. dust collection tray contents into biohazard

4. ethanol can be good for getting off stuck paraffin

mouse work

general

• if you work with multiple strains, deal with the immunocompromised first, and swap gloves when swapping between strains, and use a new holding cage

• immunocomps have to be worked with in the hood and have their own set of tools

• hold the tail no more than halfway down as this is more secure

• when scruffing, you are looking to be holding the skin down the whole of the mouse’s back. The skin should be pinched between the length of the inside of the thumb from pad to base and the side of your index finger from the first knuckle down to the join on the thumb. You press down and slide your two-fingered grasp up the mouse till you reach behind the head, then pull the skin up and taut to scruff.

• No fingers near the face - it scares them

• Grab by the tail rather than the body - they prefer this

shaving

• need a good scruff - don’t have the head grabbed and don’t have the chest pulled too tight or they won’t be able to breathe

• shave a larger area than needed to make sure the tumor is clear

• shave as close to injection day as possible

• the hair grows in all different directions so you need to shave every way

• don’t need to exert too much pressure

• shave over the holding cage to catch the majority of fur

• watch the mouse as you go and set them down if they need a bit of a break

• stay calm, and the mice will be calmer too

injections

• your needle is your first priority - you place and discard the needle before you deal with the mice

• no injections when hungry - makes for shaky hands

• want to restrain the mouse on its side so you have access to the tail veins

• can switch sides if you don’t like the side of the tail that you are on

• patience is key - if you don’t like how its looking, take a moment to reset

• put the mice under a normal desk lamp for 4 minutes prior to injecting so their veins enlarge due to the heat

• in the hand without the syringe, you use some fingers to push the tail in both directions to generate tension at the injection site

• aim the needle bevel down to help with piercing the skin - make sure the numbers on the needle are visible and facing you

• the mice tend to piss in the holding tube, which can make dealing with the later mice more difficult cus pheremones

• you know you have the vein when - you need barely any force to depress the plunger, the vein turns momentarily white as you administer the payload as the blood is displaced, and when you remove the needle, a drop of blood comes out. You can run your finger down the tail to give it a slight squeeze to check for blood.

• have some paper towel on hand to staunch the mouse bleeding

• fill the syringe an extra 3 sections or so

• hold the syringe point up and flick to remove bubbles, as these will kill the mouse - aspirate the initial excess taken up back into the source vial to clear out the bubbles, and you’re ready to go

• if you have to set it down, prop the needle on something (like the cap) so it isn’t directly on the table

• unless they have different treatments, fill the syringe with enough for all the mice on the table

• only one pair of gloves here, as you need good dexterity

• aim to the top right of the caivity for IP injections

• if you are having trouble aspirating from the mice, stop and reset

Jess western blot

auto wb

1. separation - add a gel separated by a charge difference, add fragments of molecular sample - fragments will move through the gel at speeds related to their size and separate out into bands

2. gel is transferred to a membrane to absorb the protein bands

3. stain composed of primary Ab sticking to target and secondary carrying the indicator sticking to the primary

Jess automates these steps

have to boil and denature the substrate so it moves through the gel

used previously to determine if proteins were being cleaved

tumor takedown spleen processing

1. Need one of those 6 well plates with about 5mL rpmi media in each. One of these plates is good for 3 spleens, with two wells per spleen.

2. Put one of those white filters to soak in each of the three wells in the top row.

3. Once soaked, take a spleen tube and dump the spleen into the filter in the top well. Take the plunger of a syringe, tip the flat plastic side in the lower well without the filter, and then use it to gently grind up the spleen. Avoid mashing any resulting fat if possible.

4. When well mashed, discard plunger in sharps (if any sizeable chunks of spleen are left on the plunger, gently wash it in the lower well, and discard all if they don't come off).

5. Take the filter, dip it gently in the lower well to dislodge anything from the bottom, then place in on a clean 50mL falcon tube. Pipette the solution from the top well (pipette up and down a couple times while tilting the plate and expelling the fluid near the top half to wash all spleen chunks to the bottom where they will be sucked up) and slowly through the filter. Slowest pipette boy speed, small squirts into the filter, lift the filter up after a few squirts enough to release the vacuum from the tube and allow the fluid to pass down, if any fat is deposited from the pipette move the tip to avoid squirting any solution off the obstructing fat.

6. Once filtered, pipette the media from the lower well (same sero), pipette up and down a few times, wash out the top well and up and down a few times, then put this through the filter as well. Do another wash with an additional 5mL of media taken from the media bottle (new sero) if the spleen is especially large (or just add it to the filtered stuff if you need to balance it anyway from another spleen tube).

7. Once done, take out the filter, scrape the bottom a few times on the lip of the falcon tube, then discard in sharps. Cap the tube, invert a few times to get any matter on the sides, and you're done.

dry ice flash

epindorph vial, piece pf dry ice, drop of water, chuck and leave

will go off randomly

never seen this done before - don’t recall who told me this