Core Facilities
USF Health - College of Medicine

     

Light Microscopy

Excitation and Emission of Fluorophores (pdf file)

Fluorescent Proteins (pdf file)

Sample slide preparation:
It is recommended to use #1.5 coverslip for sample preparation since most objective lenses are manufactured to be used with standard glass slides and coverslips of a certain thickness, usually 0.17 mm, which corresponds to the thickness of no. 1.5 coverslip.  
       
To prevent coverslip displacement and obtain the best images as possible, any excess amount of mounting medium should be removed after coverslip are placed. This can be easily done by placing a folded sheet of Kimwipes over the coverslip and gently tapping/pressing to squeeze out the excess medium until you get no excessive mounting medium. 
       
We also highly recommend that sample slides be sealed with nail polish or other sealing reagents. There are a couple of advantages of sealing; it helps to prevent samples from drying out, sliding of coverslip by the moving stages, embedded material movement during z-sectioning, and leaking of mounting medium and contaminating the immersion oil, which may blur your images and damage the lens. A cheap nail polish we recommend is the clear “Hard as Nails” from Sally Hansen available at grocery stores.

Preparation of glass coverslips for growing and imaging cells: 
For regular imaging, cells are plated on glass coverslip (12 mm circle or similar) inside of culture dish. The coverslips are typically pretreated with poly-L-lysine (PLL) if cells are loosely adherent to glass (polyornithine works better for some types of neurons).

Acid wash coverslips. This helps cells and polyamino acids stick to glass.
1. Heat coverslips in a loosely covered glass beaker in 1M HCl at 50-60 oC for 4-16 hours. 
2. Cool.
3. Wash coverslips extensively in dH2O, then ddH2O.
4. Rinse coverslips in ethanol and leave to dry between a folded sheet of whatman paper(dry as separate coverslips).
5.Keep in a sterile tissue culture dish (can store for a year).     

Coat with polyamino acid.
1. Coat coverslips in bulk in 10-15 ml of 1mg/ml PLL (or 500 mg/ml polyornithine), rocking or rotating for a minimum of 30 minutes in a 10 or 15 cm tissue culture dish.
2. Save the polyamino acid (can reuse 3-4 times).

3. Wash the coverslips in dH2O, then ddH2O at least 5 changes in each (free polyaminoacid is cytotoxic). 
4. Rinse coverslips in 100% ethanol and dry those to be used immediately on one end in an open tissue culture dish in a sterile incubator. 
5. When dry, add cells.
6. Dry remaining coverslips between a folded sheet of whatman paper (dry as separate coverslips).
7. Keep in a sterile tissue culture dish (can store for a year). Do step 4 before use. Can keep 10-20 ml aliquots of 1mg/ml PLL and 500 mg/ml polyornithine stocks at -20 oC. High molecular weight PLL is standard (greater than 300K), but lower molecular weight PLLs can also be tried.

    

Optional-- Coat polyaminoacid/acid washed/coverslips with matrix molecules. This helps the attachment of very poorly adherent cells (e.g. neurons), and increases the growth rate of other cell types (e.g. primary culture cells). Different extracellular matrix molecules can also change the morphology of certain cell types (e.g protrusion of lamellipodia or filopodia, flattening of cell bodies useful for microinjection- usually determined empirically). 1. Coat individual polyamino acid/acid washed/coverslips with a drop of specific matrix molecule (held by surface tension) from frozen stocks for a minimum of 30 minutes at room temperature or in a 37 oC incubator, or overnight at 4 oC.
Examples:

          --Collagen type IV/PLL/acid washed coverslips

               PC12 cells (100 mg/ml collagen), somites (2 mg/ml collagen).

          --1x matrigel/PLL/acid washed coverslips

               Primary fibroblasts, neuroblastomas, fibromas, amphibian motor neurons,

               embryonic dorsal root ganglia.

          --10 mg/ml laminin-polyornithine acid washed coverslips

               Adult dorsal root ganglia.
2. Wash 5x with calcium and magnesium free PBS, then 1x with culture media.

3. Plate cells. Coverslips must be coated fresh before plating cells. Washed, coated, coverslips can be stored for a maximum of one day in the cold room.

 

Basic Protocol for antibody labeling of cultured cells
(This is a generalized version of a typical staining process. You may need to adjust or modify each step to optimize staining of the proteins of your interest.) 

1)      Plate cells on #1.5 coverslips or chamber slides.

2)      Rinse culture media off with PBS or media without serum to remove serum proteins. Rinse means replacing liquid with the same amount of wash solution. No incubation necessary. You might want to move the coverslips to a 6 or 12 well dish for labeling for ease of solution changes.

3)      Add 4% paraformaldehyde in PBS, incubate at room temperature (RT) for 10~15 minutes.

4)      Rinse 3 times with PBS or PBST (PBS + 0.1% Triton-x100, with 0.5% BSA [or appropriate serum of animal where the secondary antibodies are raised]).

5)      Incubate in PBST for 30 minutes prior to adding primary antibody diluted in PBST.

6)      Incubate with primary antibody either for 1 hour at room temperature, or overnight at 4 oC.  You can add multiple primary antibodies together if they are from different species.

7)      Rinse 3 times with PBST.

8)      Add diluted secondary antibody (e.g. molecular probes goat anti-mouse Alexa 488 at 1:500) to the coverslips and incubate at room temp for 1 hour. You can add multiple antibodies together as long as they are directed against different species.

9)      Rinse 3 times with PBST.

10)   If you want DAPI labeling and your mounting media does not have it already, add DAPI at 90mg/ml and wait 15 minutes prior to mounting (no need for this step if you are using Vectashield plus DAPI mounting medium).

11)   Pick up coverslips with a forceps and touch them to a kimwipe to wick off excess PBT, put them face down on about 20-50 ml of a mounting medium (Vectashield or 90% glycerol in PBS).

DNA (nuclear) staining:
Nucleus can be stained with various reagents that binds to nucleic acid. Some dyes (i.e. DAPI,Hoechst, DRAQ5) bind preferentially to double-stranded DNA, whereas others interact with both DNA and RNA, which requires treatment of samples with RNase to enhance the DNA staining.

* DAPI (or Hoechst): This dye shows max. excitation at 350 nm and max emission at 470 n. It is minimally excited by 405 nm laser on confocal microscopes, but usually the signal is very intense enough to be easily visualized. It is convenient because of no need for pretreatment. Staining is normally performed after all other staining. Thus, if you are using Vectashield with DAPI mounting medium, just a drop of the medium takes care of DNA staining. 

Dilute the DAPI stock solution to 300 nM in PBS. Add this dilute solution just enough to cover cells on the coverslip preparation.  Incubate for 1–5 minutes and rinse the specimen several times in PBS. Mount the sample.

* DRAQ5: This is a highly cell permeable far red range DNA dye that exhibits max. excitation at 647 nm and max emission at 679 nm. Since it shows minimal affinity to RNA, staining does not need RNase treatment to reduce the RNA-bound background. Furthermore, it is very photostable and efficiently excited by 633 nm laser. Unlike DAPI, it can be used for live imaging for a short period of time with a low laser power.

With living cells, add 5-10 mM DRAQ5 to culture medium and staining can be done in less than 5 min at room temperature.

For fixed cells, cells can be mounted in a mounting medium containing 5 to 10 mM range or can be stained with PBS containing the dye during the final rinse step for 5-10 min followed by quick rinse in PBS.


* Propidium Iodide and cyanine dyes (TO-PRO, YO-PRO....): Since these dyes bind well both DNA and RNA, sample need to be treated wiht RNAse for nuclei staining. During the secondary antibody staining, add 5 mg/ml of RNase A to the sample. Then, during final wash step, add the nucleic acid dye at ~0.5 mM and stain for 30 min to 1 hr at room temperature. Rinse well and mount the sample. 

Emission spectra of DNA-bound cyanine dimers (from Invitrogen).

Comparison of Nuclear DNA stain for live cell imaging

Nuclear dyes

Propidium Iodide

TOTO3

TOPRO 3

Hoechst 33258

DAPI

DRAQ5

H2B-GFP

Practical  Aspects

Membrane permeant?

No

No

No

DAPI only semi- permeant

Yes

n/a

Live/fixed cell application?

No/Yes

No/Yes

No/Yes

Yes/Yes

Yes/Yes

Yes/Yes

Fixation or permeabilization needed?

Yes

Yes

Yes

No

No

n/a

Stains intact primary culture cells?

No

No

No

Yes

Yes

Can be difficult to transfect.

Time (minutes)/staining steps

60/5

60/5

60/5

5/1

5/1

n/a

Working concentration

1.5 µM

1 µM

50 µM

1.6 µM

1 µM

 

Biological Aspects

Nucleolar/RNA staining?

Yes

Yes

No

No

Very weak nucleolar staining

n/a

Mitochondrial staining?

No

No

No

No

No

No

Stoichiometric to histone 2b?

n/d

n/d

n/d

No

Yes

n/a

Phototoxic?

n/d

n/d

n/d

2-photon excitation leads to cell death.

No

n/a

UV damage risk?

n/d

n/d

n/d

Yes

No

n/a

Spectral aspects

Excitation/emission maximum (nm)

535/617

642/660

642/661

352/416

647/670

488/507

 Photobleaching

Slow (1-2 min)

Fast (20-30 sec)

Very fast (5-10 sec)

Slow (1-2 min)

No bleaching

Slow (~1 min)

2-photon excitation?

Yes

Yes

Yes

Lead to cell death

Yes

?