Solutions to have at the beginning of the semester

Students use lots of these:

• 10 mL PBS
• 10 mL serum-free-non-CO2 medium
• 10 mL HBSS
• 10 mL HBS

Students split a flask of B16s into a new flask and a coverslip-bottom dish for viewing next class

• 1 flask of B16-30 (wild type) per pair (at least 50% confluent)

• 1 flask, 1 coverslip-bottom dish per student

• 1 50 mL tube of DMEM per pair

• 1 15 mL tube of PBS per pair

• 1 0.5 mL aliquot of trypsin per pair

• 5 mL serologicals

• sterile transfer pipettes

• 70% ethanol spray bottles

• lots of gloves

• handout

Immunofluorescence Procedure Tu 9/26/23

Materials

• PBS-Tw-Az for rinsing
• Little beakers
• forceps
• enough for 4 humid chambers
  • petri dish
  • filter paper
  • parafilm
• fixed coverslip of HTC116 cells, 1 per student
• Mixed primary antibodies
  • Mouse anti-Lamin A
  • Rat anti-alpha tubulin
• Mixed secondary antibodies
  • Goat anti mouse 488 (green)
  • Goat anti rat TRITC (red)

To mix primaries:

• Lamin A:
  • Use 4 aliquots of 5 µL each
  • Add 125 µL PBS-Tw-Az-BSA to get (slightly less than) 2X strength
  • total volume: 520 µL
• Alpha Tubulin:
  • Use 3 aliquots of 4 µL each
  • Add 200 µL PBS-Tw-Az-BSA to get 2X strength
  • total volume = 600 µL
• Mix 1:1, and aliquot ~104 µL per pair. Make 10 aliquots

To mix secondaries

• Green anti-mouse:
  • 2 1µL aliquot
  • 300 µL PBS-Tw-Az-BSA
  • total volume: 602 µL
• Red anti-rat:
  • 2 5 µL aliquots
  • 260 µL PBS-Tw-Az-BSA each
  • total volume: 530 µL
• Mix 1:1, and aliquot ~106 µL per pair. Make 10 aliquots

Live Cell Staining & Imaging

Students use the dish they seeded previous Thursday.

Need "staining solution":

• Fluorobrite
• Nuc Blue
• Mitotracker Red

Kate transfects HCT116 cells with Eb1 plasmids.

Need 8-10 viewing dishes, seeded with transfected cells 1 to 2 days before viewing.

1 reaction needs ~ 10⁶ cells/mL

100 uL cells per reaction (barely 8 drops)

So do 2 reactions, each with 10⁶ cells.

2 days before, seed a 25 cm² flask with 0.5 10⁶cells.

(4 days before, seed with 0.125 10⁶ cells)

1 drop of the 100 uL reaction into its own dish.

On transfection day, divide into 2 reactions

Notes on actual protocol followed:

Friday 10/6

• Start with 2 confluent 12.5 cm² flasks
  • (~1.25 x 10⁶ cells)
  • Need 1-5 x 10⁶ cells/mL on Monday
  • Split 1/8 to allow for 3 doublings
  • Stop reaction with 1.6 mL
  • Seed 0.2 mL into each of 2 new flasks.

Monday 10/9/23

• Put all the cells from one confluent 12.5 cm² flask
  • should be confluent again
• into a new 25 cm² flask

Tuesday 10/10/23

• There should be 2.5 x 10⁶ cells
• Trypsinize
• Stop reaction with 2 mL
• split suspended cells into two microfuge tubes.
• Spin down, and proceed with 2 Mirus nucleofection reactions
• Make 2 12.5 cm² flasks

Trial run for Lamin antibodies

Fix 15 min in paraglut

1^o Ab: Rabbit anti-lamin B

• dilute 1:1000 with PBS-Tw-Az-BSA
• 2 µL antibody into 2 mL buffer
• 1 hr 37C

2^o Ab: goat anti-rabbit

• 1 hr RT incubation

Stain with Hoechst (1:10,000) in the final PBS-Tw-Az rinse

Mount with non-DAPI Fluoromount (Fisher OB100-01)

Students begin projects

Kate provides fixed coverslips (seeded 10/22, fixed 10/24/23)

Materials:

• Paraglut fix (enough for 30 wells)
• PBS to rinse 30 coverslips
• PB-TW-Azide (enough for 24 dunking beakers and 30 wells)
• 1^o antibodies
  • mouse anti-NP (nuclear pore)
  • mouse anti-LB1
  • rabbit anti-LB2 ab151735 (get one 10 µL aliquot from Drew - dilute 1/1000)
• 2^o antibodies
  • goat anti-mouse green for NP
  • goat anti-rabbit red for LB1 & LB2
• Slides
• slide labels
• DAPI & droppers
• forceps
• dunking beakers
• humid chamber materials

For "Learn your Microscope" 9/3

9 HeLa (Vikash) coverslips

Fixed in paraglut

Stained for alpha tubulin:

• rat anti tubulin (directions on tube)
• goat anti rat TRITC (1:1000)

Stained for green actin

Mounted in DAPI

Note: Red tubulin was gorgeous! Let's see if it lasts longer than the FITC.

Lamin B1 (rabbit, 1:1,000);

Lamin B2 (rabbit)

Goat anti-rabbit green

Emerin (mouse, 1:1,000) (from Drew's lab);

Goat anti-mouse red

Fixed coverslips (MEF 39 from Drew's lab)

Fix in paraformaldehyde

To make the double primaries:

Make 2x Lamin B1 (add 1 mL instead of 2 mL)

Make 2x NPC (follow label directions = 2x (should say 1:20)

Make 2x Emerin: 1:500

Mix 250 µL each

To make the double secondary:

• Make 2x goat anti rabbit green
  • goat anti rabbit green aliquots need 1194 µL to be 1X
  • make 2X by adding 597 µL PBS-Tw-Az BSA
• Make 2X goat anti mouse red:
  • aliquot says add 500 µL to make 1X
  • add 250 µL to make 2X (use 2 aliquots)
• Make 1x green + red by mixing
  • 500 µL goat anti rabbit green with
  • 500 µL goat anti mouse red

DAPI

Slides

Beakers

forceps

Humid chambers

PBS-Tw-Az

Hoods When beginning work in a hood, spray all walls and the working surface with 70% ethanol and wipe clean.

When you have finished working in a hood: please leave it empty, spray the walls and working surface with 70% ethanol and wipe, and make sure that the gas is off. Leave the fan on and turn on the UV light.

Incubators Immediately remove any contaminated dish or flask from the incubator and treat itwith 30% bleach. Wipe the shelf it was on with 70% ethanol. Notify any individuals with cells in the incubator of the contamination.

If you are using an incubator that requires CO2: Always check the water level in the bottom of the incubator, if level is low add water. You are responsible for checking the level of CO2 in the tanks, if pressure falls below 800 psi the tank is emptying. After having received instructions, connect a full tank to the incubator and order a replacement (or notify someone).

Microscopes Please make sure the microscopes are clean and covered when not in use.

Pipets Dispose of all glass Pasteur pipets in the glass only bin. Disposable plastic pipets should be placed in trash (in the plastic sleeve). Reusable pipets should be placed tip up in the pipet jar.

Supplies Label all items in the refrigerator with your name and date. Items that not labeled will be discarded.

Vacuum Pump When you are done, please empty and rinse flask with 30% bleach.

Other Keep the area clean at all times. Dishes and flasks of cells to be discarded should be treated with 30% bleach before trashing.

• Grow cells on coverslips in 6-well plates

• Remove coverslips using a fine-tipped forceps when desired cell density is reached, and put in coverslip rack, cell side facing "cells" label on holder.


• Put holder in 100 mL beaker containing ~60 mL PBS with a tiny stir bar. Rinse by stirring for ~5 minutes.

• Transfer holder to a second 100 mL beaker containing ~60 mL of fixative. Fix with stirring for ~10 minutes

• Transfer holder to a 3rd beaker containing PBS-Tw-Az. Rinse by stirring for ~5 minutes.

• Repeat with another beaker of PBS-Tw-Az.

• Stain or mount coverslips right away, or store them in the holder in a brown jar with fresh PBS-Tw-Az. They may last up to a month in the refrigerator. Longer than that, they will start to fall off the coverslip.

Thermo Scientific Napco Series 8000 DH

1-888-213-1790

www.labequipmentparts.com

• 37C
• 5% CO2
• pan of water in bottom

• Thermo Scientific Isotemp Gas Line Filter:

  • Fisher 15-497-026 (Thermo Scientific 760210) $176.54/Pack of 10

• Thermo Scientific Replacement HEPA Main Filter:

  • Fisher 15-497-022 (Thermo Scientific 760175) $60.60 each
  • Fisher 15-497-023 (Thermo Scientific 760209) $308.62/Pack of 4

• CO2 sensor for model 3598: part # 1900602 (~$400)

• Manuals under incubator in 371

Just before plating, after cold medium has been added to the trypsinized cells,

• mix 0.1 mL cell suspension with 0.1 mL 0.4% trypan blue. Live cells will not take up trypan blue. (Use a different dilution factor if cells are uncountable)

• inject 10 uL to one side of the hemocytometer (see image attached below).

• Inspect at 10x.

• Count the number of live (clear) cells in all 9 squares of the hemocyometer. (Volume = 3mm * 3 mm * 0.1 mm = 0.9 uL) (Count in 4 1mm x 1mm squares if cells are very numerous)

• Calculate the live cell concentration (# live cells * 1.11)/1uL for 9 squares,
  (# live cells * 2.5/uL for 4 squares)

• correct for dilution factor

Cells are in suspension -- they are not dead!

To make conditioned medium:

• spin cells to bottom of conical tube (be gentle!!).
• Save the supernatant in a labeled tube that has the date and is labeled “conditioned medium”.
• Store extra in fridge.

To grow cells:

• Grow cells at room temperature, no CO2.

• Passage every 3-4 days. Dilute cells 1:5 each split and use 20% conditioned medium.

• Put cells from flask in tube; spin; take off and save supernatant (this is your conditioned medium!)

• Resuspend pellet (gently) in 5 mL medium.
• Add to clean flask
  • 1 mL conditioned medium (supernatant)
  • 1 mL cell suspension
  • 3 mL fresh medium
• To induce gene expression (e.g, tubulin) add copper sulfate to the medium
  • cells are neomycin resistant
  • Induce 1 day prior to imaging
  • to image put on previously prepared ConA coated coverslips. Image 30 min after plating on Con-A coated dishes or Con-A coated coverslips

To make medium for growth of Drosphila cells:

Schneider's medium + 10% heat-inactivated FCS.

Freezing S2 cells

Spin down a 3-4 day old 75 cm² confluent flask. 1200 RPM, 5 min

Remove conditioned medium

Resuspend one flask's worth of cell pellet in "freezing medium"

• 1.8 mL fresh medium (including antibiotic and serum)
• 1.8 conditioned
• 400 uL DMSO

Make 1mL aliquots.

Put in -80 in styrofoam for 1 day.

Transfer to liquid nitrogen.

Approximately 1 freezing vial per 25 cm2 flask.

3 vials from large size (75 cm2) culture flask.

Feeding Cells before freezing

• When cells are 90% confluent (~4 days), replace medium
• Draw out old medium, replace with 5 mL new (warm) medium.

Freezing cells

• Make freezing medium (15% DMSO, 20% serum) in advance
• Remove medium from flask
• Rinse once with ~3 mL sterile PBS. (Squirt in, then suck out.)
• Add 10 – 15 drops trypsin (0.5 – 0.75 mL)
• Incubate (37C) 5 – 30 min
• Add 3 mL cold medium (regular)
• Pour into 15 mL centrifuge tube
• Optional – rinse with another 2 mL, to capture as many cells as possible
• Spin (~100-500xg) 5 – 10 min (until supernate is clear and pellet is hard)
• Pour off supernate (if pellet is solid – otherwise pipette it off)
• Resuspend in 1 mL freezing medium (pipette up and down)
• Transfer to freezing vial (should be slightly more than 1 mL)
• Put on ice
• Store overnight in -80C in "Mr. Frosty" (actually a VWR 414004-284 CryoCooler Freeze Controller)
• Put into liquid nitrogen.

Fill out the freezing log

Frozen cell lines

• Put medium into dishes
  • MatTek dishes for observations of live cells
  • coverslip in a 35 mm dish for fixation and mounting
• Let dishes equilibrate in incubator
• Treat cells in flask with trypsin as for splitting
• Stop trypsinization with cold medium
• Add 1 - 4 drops of medium + cells to each dish, depending on cell density
• Check density of first dish, adjust volume for the remaining dishes
• Check after 24 hours.

• Put 5 mL medium in each flask. 2.5 ml for mini flasks. Label with cell type and passage number
• Warm flasks in incubator
• Remove medium from cells.
• Right away, Rinse once with ~3 mL sterile PBS. (Squirt gently in, Rock the flask, and then suck out. Do not hit the place where cells are. )
• Add 15 drops sterile trypsin (=~0.75 mL) - enough to cover cells, if more than a thin layer, pull some out
• Incubate (37C) ~5 min or until all floating, likely no more than 5m.
• Check to make sure you can see them floating in the liquid
• Add ~1mL medium to flask, draw up and down to mix.¹
  • ~4-8 drops (= 0.2 – 0.4 mL) into each new flask containing medium ²³
  • ~4-5 drops ( = 0.2 – 0.25 mL) if cells are 70 – 80% confluent
• Put back in incubator.

counting

• Label flask with cell line, passage number, date
• Put 10 mL medium in flask, set in incubator
• Retrieve frozen vial from LN2 dewar, thaw in incubator or water bath.
• Put ~1 mL thawed cells into a single flask
• 24 hours later, replace medium (5 mL)
• Split as needed

need dapi slides - can use the same ones for 2.2

squuze bottle of ethanol to clean oil off lenses.

Stage micrometer:

10X: 2.162 pixels/µm

40X: 8.728 pixels/µm

Reticle is 2 mm

Major divisions are 100 µm

tiniest divisions are 10 µm

Fluorescence Microscopy

LLCPK fixed and stained with DAPI.

Prepare before the semester starts, keep in a slide box in the refrigerator.

Bring to room temperature in the morning of the lab to minimize condensation on coverslip.

Keep track of which number slides have been used, so they will not be used later in any lab where photobleaching is an issue.

Immunofluorescence Labeling

Turn on the Incubator

Reagents

• PBS (warm) for rinsing medium off cells before fixation

• PBS-Tw-Az for rinsing coverslips (in carboy)

• 2% BSA in PBS to make antibodies

• Fixative

  • paraglut for students
  • methanol in freezer for certain antibodies

• Slides

• Nail polish

Mystery Antibodies for 2012

Direct Fluorescent Labeling of Organelles

CHECK THIS FOR HBS vs PBS!!

Cells

3t3, LLCPk

• Fixed in paraglut for phalloidin
  • 8 3t3 (can be fixed as late as the morning of lab)
  • 8 LLCPk (can be from pre-semester prep)
• Live on small diameter MatTek dishes for Mitotracker and ceramide
  • 8 3t3
  • 8 LLCPk

Reagents

• PBS-Tw-Azide

• HBS to incubate ceramide

Live cells on MatTek dishes:

Task 2. GFP-alpha-tubulin cells (everybody) 8 dishes

Task 3. Nucleofected with GFP version of (2 different ones per group) 4 - 8 dishes each:

• Rab11
• E cadherin
• H2B (GFP)
• alpha actinin

Cell Cycle

Fixed and mounted coverslips of LLCPk cells, stained with DAPI and anti-tubulin

• normal

• serum starved

Cells won't adhere to the coverslip without serum, so grow them first for 24 hours in regular medium.

Remove the medium, replace with serum-free medium

Fix the coverslips 24 hours later.

• Nocodazole treated

Motility

Live cells on MatTek dishes

8 MatTek dishes each:

• 3t3

• B16 (do more than one set, at different concentrations)

• LLCPK actin

Must be plated lightly for motility.

Scheduling

• Thaw LLCPK-actin, B16 24 hours before plating

• Plate on all cells (lightly) Monday for Wednesday lab

3t3’s must have 36-48 hours to stretch out.

If they get overgrown (especially B16s), scrape some off

Reagents

Aliquot non-CO2, serum-free medium

CHECK CYTOCHALASIN STOCKS FOR 2012

2012: 500 nM (250 nM final in dish) should give an effect without causing massive detachment of cells

Stock is 10 mM in DMSO.

Need 6 1 mL aliquots of 500 nM in non-CO2 medium:

• 1 µL 10 mM stock
• 20 mL medium

2011: working concentration of cytochalasin D: 1-10uM

Make 200uM stock for students to dilute with non-CO2 medium

4 groups using cytochalasin, each doing 4 dishes.

Maximum conc 10 µM, in 2 mL

vi = (10 µM * 2 mL)/200 µM = 0.1 mL

So each group would use at most 0.4mL

Stock was 10 µL of 10 mM cytochalasin D in DMSO.

Adding 450 µL of non-CO2 medium makes it 500 µL of 200 µM
cf = (10 mM * 10 µL)/500 µL = 0.2 mM = 200 µM

Frozen stock is 2.5 mM
Invitrogen PHZ1063

Make 500 µL of 0.2mM
vi = (0.2mM * 500 µL)/2.5 mM = 40 µL
Add 460 µL non-CO2 medium

40 µL aliquots. Each can make 500 µL of 200 µM for use.

Nocodazole

Taxol

Receptor-mediated Endocytosis

Monday 10/17/11

Bulk-phase Endocytosis

Transport of Endosomes along Microtubules

Wednesday 10/26/11

Release of Internal Calcium Stores

Wednesday, 11/2/11

Rescheduled for Monday, 11/7/11, due to snowday 10/31/11

Projects

Have students make a class poster for the bulletin board in the hall.

For Wed, 11/14/12

Plating:

• LL parental: 5 coverslips, 7 dishes
• LL alphas: 4 dishes, 2 coverslips
• LL alpha, myo: 4 dishes
• 3t3: 4 dishes

W 9/04

Intro to Equipment

W 9/09

Imaging mitosis with fixed cells. Image Spatial Quantification

W 9/11

Image Formation in the Microscope

M 9/16

Numerical Aperture

W 9/18

Resolution

M 9/23

Fluorescence Microscopy

W 9/25

Identification of Cellular Organelles

M 9/30

Labeling Cellular Structures

Cells

• paraglut fixed LL's for students to stain for tubulin (make extra!)

• paraglut fixed LL's for students to stain for phalloidin

• paraglut fixed 3t3's for students to stain for phalloidin

• live LL's to fix in paraglut (2014: treat first with mitotracker, then fix in formaldehyde no detergent)

Solutions

• PBS to rinse coverslips

• paraglut

• formaldehyde

• PBS-Tw-Az

• 2% PBS-BSA to make up antibodies

Materials

• forceps

• 50 mL beakers (switch to holders and 100 mL beakers?)

• petri dishes for humid chambers

• filter paper for humid chambers

• mounting medium

• slides

• nail polish

• kimwipes

• parafilm

W 10/2

Imaging Living Cells Expressing GFP-Tagged Proteins

• LL-GFP-tubulin

• LL-GFP-actin

• LL nucleofected with plasmids

  • H2B

2014

• LL-EB1-GFP stable line

• LL-alpha tubulin=GFP stable line

** transfected**

• H2B m cherry

• alpha actininin (GFP?)

W 10/9

Motility in Control Cells

On MatTek dishes:

• 3t3

• B16

• LL-GFP actin

Non-CO2 medium

Tu 10/15 & W 10/16

Mechanism of Motility

Students check for effect of dose and time on actin filaments. Treat LL parentals with cytochalasin D, then stain actin with phalloidin.

• LL on coverslips

• non-CO2 medium

• Cytochalasin D

• paraglut

• PBS-Tw-Az

W 10/23

Receptor-mediated endocytosis

NO LL's

NO high iron

3 coverslips of 3t3 per group

Try Lysotracker (Invitrogen L-7528)

• 50 - 75 nM

• stock = 1 mM in DMSO

• 30 min - 2 hours incubation warm.

• Add lysotracker to cells at 1 pm.

  • Make 100 µM stock
  • 1 µL of 100 µM stock to each dish. Assume ~1.5 mL per dish

Omit the high-iron treatment

For convenience, all staining, incubation, etc. will be done in HBS.

Each group does 3 coverslips of 3t3.

• HBS to mix everything else in. There is a bottle of sterile 10X on the prep room bench.

  • 6 mL aliquots COLD (final rinse before fix coverslips 1-3)
  • 15 mL aliquots RT in hood (to rinse off fixative)
  • Need ~300 mL total

• Fe-HBS

  • 20 mL aliquots COLD to rinse off growth medium, then tf
  • 5 mL aliquots WARM (incubation after tf treatment)
  • need some to make Fe-HBS-BSA for transferrin
  • make 200 mL

plus water to final volume

• Fe-HBS-BSA 1 mg/mL to make transferrin solution

  • 5 mg BSA in 5 mL Fe-HBS

• Transferrin in Fe-HBS-BSA

  • Alexa-Fluor 488 transferrin (Invitrogen T13342, MW = ~80KD) stock is 5 mg/ml (=62.5 µM). Final concentration = 1 µL.
  • Each group does 3 50-µL treatments (need extra for pipetting error)
  • Make 1.2 mL: 19.2 µL AF-488-Tf + 1.1808 mL Fe-HBS-BSA
  • Aliquot 165 µL
    if new transferrin stock is needed add 1 mL sterile distilled water to the new bottle.

• HBS-paraformaldehyde 3.7%

  • Need 60 mL
  • Divided in 2 aliquots, one per fume hood

Old fashioned way:

heat paraformaldehyde in water, then add 10X HBS
Add a drop of 10N NaOH
Heat and stir in the fume hood

OMIT HIGH IRON

plus water to final volume

M 10/28

Bulk-phase endocytosis

** 2 coverslips, 2 small diameter MatTek dishes 3t3

PBS for rinsing live cells WARM
PBS for rinsing fixed cells RT
Take from the supply of sterile aliquots

Growth media for incubation WARM
10 mL aliquots each

• DMEM for 3t3 cells
• non-CO2 medium for observing live cells (could use this for all cells for simplicity's sake)

TMR-dextran to stain endosomes in cells on coverslips
Invitrogen D3308, 10 mg
Make 10 mg/mL stock (add 1 mL to the bottle)

Make 8.8 µL aliquots in 2 mL tubes.

TMR-dextran Working concentration = 0.05 mg/mL in medium
2 coverslips at 50 µL/coverslip (=~110 µL/group)

Add 1751.2 µL medium to 8.8 µL 10mg/mL stock to make 0.05 mg/mL working solution.
Aliquot ~105 µL/group

TMR-dextran/ FITC-dextran to stain endosomes in cells in MatTeks
tmr as above

FITC-dextran Working concentration = 0.5 mg/mL in medium
(1mg/mL might be better, but that uses up a whole bottle.)
2 dishes at 100 µL/dish (=~220 µL/group)

Add 1742.4 µL medium to an 8.8 µL aliquot of 10 mg/mL TMR-dex plus 8.8 µL 100 mg/mL FITC-Dex

Methylamine solution (1M)
Sigma 426466-100mL 40% w/vol (=12.9M; MW=31) ~$25
(Working concentration ~20mM - ~20µL/mL. Students need ~100 µL)

Make 1 mL, aliquot ~120 µL

plus water to final volume

Fixative

• PBS-paraformaldehyde 3.7%
  
  • Need 70 mL
  • Divided in 2 aliquots, one per fume hood

heat paraformaldehyde in water, then add 10X HBS
Add a drop of 10N NaOH
Heat and stir in the fume hood

OR http://wahoo.nsm.umass.edu/content/paraformaldehyde-fix

Cells